0 THE TEXAS JOURNAL OF SCIENCE PUBLISHED QUARTERLY BY THE TEXAS ACADEMY OF SCIENCE Volume 60 Number 1 February 2008 GENERAL INFORMATION MEMBERSHIP. -Any person or member of any group engaged in scientific work or interested in the promotion of science is eligible for membership in The Texas Academy of Science. For more informa¬ tion regarding membership, student awards, section chairs and vice¬ chairs, the annual March meeting and author instructions, please ac¬ cess the Academy’s homepage at: www.texasacademyofscience.org Dues for regular members are $30.00 annually; supporting mem¬ bers, $60.00; sustaining members, $100.00; patron members, $150.00; associate (student) members, $15.00; family members, $35.00; affili¬ ate members, $5.00; emeritus members, $10.00; corporate members, $250.00 annually. Library subscription rate is $50.00 annually. The Texas Journal of Science is a quarterly publication of The Texas Academy of Science and is sent to most members and all sub¬ scribers. 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No part of this publication may be reproduced, stored in a retrieval system or transmitted, in any form or by any means, electronic, mechanical, recording or otherwise, without the prior permission of the Managing Editor of the Texas Journal of Science. THE TEXAS JOURNAL OF SCIENCE Volume 60, No. 1 February, 2008 CONTENTS Trophic Status Related to Periphyton and Macrophyte Distribution and Abundance along the North Bosque River in North Central Texas. By Anne M.S. McFarland, Jimmy S. Millican, Jeffrey A. Back and Jeffrey S. Brister.................................................................................. ........... 3 Distribution of the Bantam Sunfish, Lepomis symmetricus (Perciformes: Centrarchidae), in Arkansas. By Chris T. McAllister, Renn Tumlison and Henry W. Robison........ . . . 23 A Positive Control for Detecting Heteroduplexes in DGGE for Microbial Community Fingerprinting. By Andrew Syvyk, Armen Nalian, Michael Hume and Alexandra Martynova-VanKley . . . . . . 33 Survivorship of Radio-Transmittered Urban White-Winged Doves on the Edwards Plateau, Texas. By Michael F. Small, John T. Baccus and T. Wayne Schwertner . . . ....45 General Notes Some Observations on the Mating Behavior of the Giant Walking Stick, Megaphasma dentricus (Orthoptera: Phasmidae). By Tara L Maginnis, Christina L Cool and Jana L Muniz . . . 57 First Report of Ochetosoma aniarum (Digenea: Ochetosomatidae) from the Brazos Water Snake, Nerodia harteri (Serpentes: Colubridae), in Texas, with a Summary of Definitive Hosts of this Parasite. By Chris T. McAllister and Charles R. Bursey . 63 Introduction of the Bluefm Killifish (Lucania goodei) in Texas. By Benny J. Callaway, Robert G. Fechhelm and Robert G. Howells . . .....69 Author Instructions . . . . . . . . . . 73 Membership Application 80 THE TEXAS JOURNAL OF SCIENCE EDITORIAL STAFF Managing Editor: Ned E. Strenth, Angelo State University Manuscript Editor: Frederick B. Stangl, Jr., Midwestern State University Associate Editor for Botany: Janis K. Bush, The University of Texas at San Antonio Associate Editor for Chemistry: John R. Villarreal, The University of Texas-Pan American Associate Editor for Computer Science: Nelson Passos, Midwestern State University Associate Editor for Environmental Science: Thomas LaPoint, University of North Texas Associate Editor for Geology: Ernest L. Lundelius, University of Texas at Austin Associate Editor for Mathematics and Statistics: ^ E. Donice McCune, Stephen F. Austin State University Associate Editor for Physics: Charles W. Myles, Texas Tech University Manuscripts intended for publication in the Journal should be submitted in TRIPLICATE to: Dr. Frederick B. Stangl, Jr. TJS Manuscript Editor Department of Biology Midwestern State University Wichita Falls, Texas 76308 frederick.stangl@mwsu.edu Scholarly papers reporting original research results in any field of science, technology or science education will be considered for publication in The Texas Journal of Science. Instructions to authors are published one or more times each year in the Journal on a space-available basis, and also are available on the Academy's homepage at: www.texasacademyofscience.org AFFILIATED ORGANIZATIONS American Association for the Advancement of Science, Texas Council of Elementary Science Texas Section, American Association of Physics Teachers Texas Section, Mathematical Association of America Texas Section, National Association of Geology Teachers Texas Society of Mammalogists TEXAS J. OF SCI. 60(l):3-22 FEBRUARY, 2008 TROPHIC STATUS RELATED TO PERIPHYTON AND MACROPHYTE DISTRIBUTION AND ABUNDANCE ALONG THE NORTH BOSQUE RIVER IN NORTH CENTRAL TEXAS Anne M.S. McFarland, Jimmy S. Millican, Jeffrey A. Back and Jeffrey S. Brister Texas Institute for Applied Environmental Research Tarleton State University, Mail Stop T04J0 Stephenville, Texas 76402 Abstract-A two-year survey was conducted to estimate periphytic algae and macrophyte diversity and abundance at 1 1 locations within the North Bosque River watershed. Ash free dry weight by species was used to measure biomass. A total of 30 different species were collected; 24 macrophytes, five periphyton, and one moss. Macrophytes comprised the largest portion of total biomass at most sites, while algae were most frequently collected. The most biomass was contributed by the macrophyte Justicia americana (L.) Vahl, while Cladophora sp. was the dominant algae. The majority of algae were found in riffles, while macrophyte biomass was fairly evenly distributed between pool and riffle habitats. Nutrient concentrations generally showed saturated conditions for periphyton growth indicating that factors other than nutrients, such as light, may be limiting periphyton and possibly macrophyte biomass along the river. In comparison to proposed trophic state indices for periphyton, mesotrophic or eutrophic conditions were indicated at four of 1 1 sites. In contrast, measurements of instream chlorophyll-a concentrations of phytoplankton indicated mesotrophic conditions at eight of 1 1 sites. These differences in tropic status based on periphyton and phytoplankton indicate a need to monitor attached, as well as free-floating algae, when evaluating for water quality impairment in small to intermittent sized creeks and rivers. In 1998 the North Bosque River was placed on the Texas Clean Water Act section 303(d) list as an impaired water body due to excessive nutrients and plant growth under narrative water quality standards (TNRCC 1998). Besides narrative standards, in Texas, excessive algal growth may be identified through violation of water quality standards for dissolved oxygen (TCEQ 2003). Concern regarding eutrophication may be defined by comparing nitrogen, phosphorus, or sestonic chlorophyll-a (CHLA) concentrations to screening levels. These screening levels are based on an 85- percentile ranking of State water quality data for specific water body types (TCEQ 2003). This ranking allows a relative indication of water quality concerns for nutrients and planktonic algae that is statistically derived but has no direct biological meaning. 4 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 For the North Bosque River, phosphorus was identified as the nutrient limiting algal growth (Kiesling et al. 2001). Exeessive nutrients, partieularly phosphorus (P) in freshwater systems, are major factors in causing excessive algal growth (Gibson 1997). To help control algal growth, two total maximum daily loads (TMDLs) were approved with a reduction goal of about 50 percent in soluble reactive phosphorus along the North Bosque River (TNRCC 2001). Routine monitoring, as part of the post-TMDL implementation effort, includes measurement of sestonic CHLA, which primarily represents the phytoplankton or free-floating algae in the water column. Sestonic CHLA concentrations are often used to measure trophic status as an indicator of eutrophication in lakes and reservoirs (Carlson 1977). Trophic boundaries proposed by EPA (2000) for phytoplankton in rivers and streams are 10 pg/L CHLA for the oligotrophic to mesotrophic boundary and 30 pg/L for the mesotrophic to eutrophic boundary. Although phytoplankton is a good indicator of eutrophication, in most cases phytoplankton measurements alone underestimate algal biomass in wadeable streams. Periphyton, as attached filamentous algae, is often dominant relative to phytoplankton (Allen 1995). Monitoring periphyton biomass is much more time consuming and labor intensive than measuring sestonic algae, so much less research has been done on defining boundaries for trophic classifi¬ cation of periphytic algae. Despite the limited research, EPA (2000) proposes boundaries of 20 mg/m^ CHLA for oligotrophic to mesotrophic and 70 mg/m^ CHLA for mesotrophic to eutrophic for mean periphytic algae. The mesotrophic to eutrophic boundary is relatively close to the 100 mg/m^ CHLA proposed by Welch et al. (1988) for defining nuisance levels of periphytic algae. An overabundance of macrophytes is also recognized as a potential water quality problem that is often overlooked in routine water quality assessments (Newton & Jarrell 1999). Macrophytes have clogged intakes for irrigation systems and can interfere with boating and fishing activities (Sosiak 2002). No current trophic MCFARLAND ET AL. 5 guidelines exist for macrophytes, although management of macro¬ phytes is also important to maintaining healthy aquatic ecosystems (Chambers et al. 1999). While instream planktonic CHLA concentrations are often monitored with regard to eutrophication, the excessive growth of periphyton and macrophytes are rarely considered with routine water quality monitoring, but may be just as important in defining impairment. The purpose of this study was to estimate the diversity and abundance (as biomass) of macrophytes and periphyton along the North Bosque River and its major tributaries and associate this information with measures of trophic status as indicators of stream eutrophication. Study Area The North Bosque River is a fifth-order stream located in north central Texas that begins in Erath County and flows southeast for nearly 179 kilometers through Hamilton, Bosque, and McLennan counties until it enters Lake Waco near Waco, Texas (Figure 1). The mainstem of the North Bosque River for the most part is cut down to bedrock with permanent flow occurring primarily in the lower reaches and intermittent flow in the upper and mid-reaches. A small portion of the upper reach maintains a permanent flow due to discharge from the Stephenville wastewater treatment plant (WWTP). The flow associated with contributions from the Stephenville WWTP can dissipate within a few kilometers, particularly in the summer when rainfall-runoff events are infrequent and generally small and evaporation is at its peak. Lower reaches of the North Bosque River maintain some base flow most notably from seepage through fractured limestone along stream banks in the lower portion of the watershed. A relatively small contribution to base flow is from municipal WWTPs along the river. The sampling sites represented a broad range of conditions within the watershed (Table 1). Sites were chosen to characterize 6 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Figure 1. Location of sampling sites used for macrophyte and periphyton survey. the flora present along the North Bosque River and in the lower reaches of several major tributaries. Included were seven sites along the North Bosque River and four major tributaries (Figure 1). Study reaches at each site were selected that had both riffle and pool habitats, were easily accessible, and were expected to have water present throughout much of the year. MCFARLAND ET AL. 7 Table 1 . Land use and drainage area characteristics above sampling sites. Site Wood/ Range (%) Pasture (%) Cropland (%) Dairy Waste Application Fields (%) Urban (%) Other (%) Drainage Area Hectares Near WWTP Influence BO020 48.5 29.6 6.6 13.1 1.5 0.7 21,700 No BO040 47.3 28.4 7.0 12.3 4.4 0.7 25,700 Yes BO060 57.4 22.8 6.0 9.9 3.3 0.5 48,900 No BO070 65.3 18.9 6.1 7.3 2.0 0.5 93,100 No BO080 68.4 16.3 7.0 6.3 1.7 0.3 146,000 No BO085 71.0 14.7 7.6 4.9 1.6 0.3 190,000 No BO090 71.5 14.0 9.1 3.7 1.5 0.2 253,000 No DC060 74.4 7.6 8.7 8.7 0.6 0.0 23,200 No GClOO 67.4 18.1 7.6 5.8 0.6 0.4 25,200 No MC055 73.6 12.5 13.6 0.0 0.3 0.0 47,200 No NC060 75.8 10.7 13.2 0.0 0.2 0.0 35,300 No Materials and Methods Sampling was conducted in March, May, July, August, and October 2004; and January, April, and June 2005 for a total of eight events. Macrophytes were identified to species and periphytic algae to genus in most cases. Diatoms were identified only to Bacillariophyta. A few unknowns were documented that could not be reliably identified. Abundance was measured quanti-tatively by obtaining the ash free dry weight (AFDW) of each species of macrophyte and periphyton collected. Quantitative sampling of macrophytes and periphyton within pool and riffle habitats along each reach was accomplished using a Surber sampler as a 0.093 m^ plot frame. The lengths of both habitat types ranged from 30 m to 50 m for riffle habitats and 50 m to 60 m for pool habitats. Sampling locations within the stream were determined using a two number coordinate system consisting of points parallel and perpendicular to the length of the stream. A list of random numbers selected prior to sampling was used to reduce the sampler’s bias in determining sampling coordinates. A total of 12 sampling points were determined for each site with six sampling points within each of the two habitat types (pool and riffle). Of the six sampling locations within the riffle, a minimum of two were in the margin areas, while the other four could be collected from the margins or THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 within the channel. Only margin samples were collected within the pool habitat, because greater water depth and turbidity in the channel portions of pools greatly limited the growth of macrophytes and periphyton. An additional reason for sampling only the margin areas of pools was to avoid collecting periphyton that may have been detached through scouring and deposited into the deeper channel portions of the pool. Macrophytes and periphyton were identified in the field when possible and recorded on the datasheet. Samples were placed in labeled plastic bags and stored on ice for transportation back to the lab. A reference collection of macrophytes collected nearby but outside the study reaches was maintained. Macrophytes used for the reference collection were dried, mounted, and labeled and stored in the Tarleton State University herbarium. Because periphyton are so fragile when dried, a reference collection of periphyton was not maintained, but any unknowns were preserved in water until they could be identified later in the lab. Because many periphyton are very difficult to identify reliably to the level of species (Prescott 1954; Dodds & Gudder 1992), all periphytic algae specimens were identified only to the level of genus, except diatoms, which were identified only to Bacillariophyta. For macrophytes, specimens were identified to the level of species when possible. All weights were obtained using an analytical balance and recorded to the nearest milligram. Samples were placed into a drying oven for a minimum of 24 hours at 105°C and weighed to obtain a dry weight. Dried samples were placed in a muffle furnace for a minimum of 6 hours at 550°C and weighed to obtain an ash weight (Barber et al. 1999). After obtaining a dry weight and an ash weight, the ash-free dry weight (AFDW), which represents the amount of organic material (biomass) present, was calculated as follows: AFDW = (Dry Weight - Tray Weight) - (Ash Weight - Tray Weight) MCFARLAND ET AL. 9 Table 2. Suggested stream trophic status boundaries for periphyton and phytoplankton based on chlorophyll (ERA 2000) and estimated boundaries for periphyton based on conversion of AFDW using equations provided by Biggs (2000). Oligotrophic- Mesotrophic- Variable (units) Mesotropic Eutrophic Boundary Boundary Mean Periphyton Chlorophyll (mg/m^) 20 70 Maximum Periphyton Chlorophyll (mg/m^) Mean Periphyton AFDW (g/m^) 60 200 7 13 Maximum Periphyton AFDW (gW) 12 24 The total AFDW calculated as the sum AFDW across all eight sampling events by species was used as an indicator of species dominance. The total AFDW by site was calculated separately for the categories of periphyton and macrophyte across riffle and pool habitats and by habitat type to indicate abundance of biomass at a site. To compare biomass abundance between sites by plant category (periphyton or macrophyte), the analysis of variance and least square means procedures were used on the total AFDW by site and sampling date to determine if significant differences (a = 0.05) existed between sites across sampling periods and to indicate which sites were different from one another. The analysis of variance and least square means tests were conducted using the SAS procedure PROC GLM (SAS 2000). A log transformation of the AFDW results was required to obtain a more normally distributed data set for comparisons between sites prior to analysis. The SAS pro¬ cedure PROC UNIVARIATE was used to confirm the need for a natural log transformation of the data using the Shapiro- Wilks statistic as a test for normality (SAS 2000). Both CHLA and AFDW are common measures of periphytic algal abundance, but developing relationships between these two measurements to approximate one from the other is a very tedious task. To compare the AFDW measurements to trophic boundaries based on CHLA proposed by EPA (2000) for periphyton, the following equations developed by Biggs (2000) were used: 10 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Table 3. Summary of marcophyte and periphyton species encountered ranked by the number of times collected (N) out of 1068 total sampling points. Species Plant Type Total AFDW (g/m") Mean AFDW (g/m") N Cladophora sp. Periphyton 3,390 9.53 356 Justicia Americana (L.) Vahl Macrophyte 14,900 97.5 153 Spirogyra sp. Periphyton 347 4.45 78 Najas guadalupensis (Spreng.) Magnus Macrophyte 83.6 1.37 61 Char a sp. Periphyton 89.9 1.58 57 Car ex sp. Macrophyte 2,000 43.6 46 Hydrocotyle umbellate L. Macrophyte 411 10.3 40 Eleocharis sp. Macrophyte 545 24.8 22 Diatom mat Periphyton 339 24.2 14 Marsilea vestita Hook. & Grev. Macrophyte 20.7 2.59 8 Ludwigia peploides (Kunth) Raven Macrophyte 96.8 16.1 6 Unidentified Macrophyte #3 Macrophyte 93.4 15.6 6 Unidentifed Algae mixed with Diatoms Periphyton 80.6 13.4 6 Selaginella apoda (L.) Spring Moss 43.2 7.20 6 Diatom mat with Spirogyra Periphyton 13.8 2.29 6 Diatom/ Cladophora mix Periphyton 26.2 5.24 5 Cyperus acuminatus Torr. & Hook. Ex Torr. Macrophyte 37.9 9.48 4 Ranunculus sceleratus L. Macrophyte 11.9 2.98 4 Poa sp. Macrophyte 51.3 17.1 3 Unidentified Macrophyte #4 Macrophyte 29.1 9.69 3 Spirogyra sp., Chara sp., Najas sp. mix Periphyton and Macrophyte 17.3 5.77 3 Zizaniopsis miliacea (Michx.) Doell & Aschers. Macrophyte 274 137 2 Paspalum distichum L. Macrophyte 90.3 45.2 2 Undientified Macrophyte #1 Macrophyte 73.5 36.8 2 Chaerophyllum tainturieri Hook. Macrophyte 12.4 6.19 2 Rorippa sessiliflora (Nutt.) A.S. Hitchc. Macrophyte 6.46 3.23 2 Cynodon dactylon (L.) Pers. Macrophyte 6.06 3.03 2 Rorippa nasturtium-aquaticum (L.) Hayek Macrophyte 1.40 0.70 2 Veronica peregrina spp. Xalapensis (Kunth) Pennell Macrophyte 1.08 0.54 2 Steigoclonium sp. Periphyton 0.32 0.16 2 Panicum dichotomiflorum Michx. Macrophyte 6.99 6.99 1 Ceratophyllum demersum L. Macrophyte 0.75 0.75 1 Unidentifed Macrophyte #2 Macrophyte 0.54 0.54 1 Smilax sp. Macrophyte 0.22 0.22 1 MCFARLAND ET AL. 11 Table 4. Sites ranked by total AFDW summed for all eight monitoring periods. Site Total AFDW (g/mO Total Periphyton AFDW (g/m2) Total Macrophyte AFDW (g/m2) Number of Periphyton Species Number of Macrophyte Species GCIOO 12,300 975 11,325 4 11 DC060 4,410 945 3,465 4 10 BO070 1,340 811 529 3 7 BO090 1,020 207 817 4 4 BO080 899 440 459 3 8 NC060 865 82 783 3 3 BO040 763 222 541 2 4 BO020 478 66 412 2 2 BO085 227 168 59 3 8 MC055 201 81 120 4 3 BO060 196 21 175 2 4 ln(CHLA, mgW) = 0.338+ 1 .396*ln(AFDW, g/m^) = 0.79 ln(AFDW, gW) = 0. 1 86 + 0.566*ln(CHLA, mg/m^) = 0.79 These equations provided by Biggs (2000) were based on detailed work conducted on 170 samples collected from a wide range of periphyton communities in streams throughout New Zealand. Trophic boundary values (Table 2) of AFDW as mean and maximum values were then compared to mean AFDW concen¬ trations across sampling events for the North Bosque River sites. Sestonic CHLA collected as part of routine monitoring along the North Bosque River was also compared to trophic boundaries for phytoplankton. Grab samples for sestonic CHLA were analyzed using a spectrophotometer following Standard Method 10200H (APHA 1992), Results A total of 30 different species were collected; 24 macrophytes, five periphyton, and one moss (Table 3). Macrophytes comprised the largest portion of the total biomass, while periphyton species were most frequently collected at all sites except GCIOO. The 12 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 majority of biomass obtained, except at sites BO070 and BO085, came from macrophytes rather than periphyton. The species most often encountered was the alga Cladophora sp. followed by the macrophyte Justicia americana (L.) Vahl (American water- willow). The amount of biomass collected at each site across habitat types was compared two ways. One by ranking the total biomass collected across all sampling periods by site, and two by statistically comparing mean biomass by site across sampling dates. The largest amount of total biomass collected at the 1 1 sites came from GClOO as well as the highest number of different species encountered (Table 4). Comparing between sites using analysis of variance (ANOVA) and least square means (LSD) procedures, significant differences (a = 0.05) in periphytic algal biomass (Figure 2a) and highly significant differences (a = 0.01) in macrophyte biomass (Figure 2b) were found between sites. The geometric mean of biomass obtained from periphyton species ranged from 0.03 g/m^ at site BO060 to 3.32 gW at site DC060 (Figure 2a). The geometric mean biomass obtained from macro- phyte species ranged from 0.01 g/m at site BO085 to 22.1 g/m at site DC060 (Figure 2b). Although the largest amount of total biomass was indicated at GClOO, the largest amount of biomass across sampling periods most consistently occurred at DC060. Macrophytes comprised 74% of the total biomass at DC060 and 92% at GClOO for the nine sampling periods, and the dominant species at both sites was the macrophyte J. americana. Justicia americana was one of the largest macrophytes collected. The erect stems of J. americana can reach up to 1 00 cm tall, and this plant is known for its extensive adventitious rooting system that can firmly bind to the streambed (Diggs et al. 1999). The abundance of J. americana at DC060 and GClOO may be attributed in part to the physical characteristics of these reaches. Justicia americana is sensitive to shading or light limitation (Fritz et al. 2004). Light availability is considered more important than MCFARLAND ET AL. 13 Figure 2. Geometric mean AFDW from (a) periphyton and (b) macrophytes for each site across all sampling events. Means followed by the same letter were not significantly different at a probability level of 0.05 according to a test of least significant differences. flooding disturbance or parasite abundance as a factor affecting total biomass of J. americana as evaluated for six south-central Alabama streams (Fritz et al. 2004). In earlier work by Howell (1975) cited by Fritz et al. (2004), the coverage of J. americana in a Kentucky stream was more than twice as much in reaches flowing east to west than in reaches flowing north to south. This greater coverage was attributed to longer daily exposure to direct sunlight of east-west reaches compared to north-south reaches. 14 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 The reach at DC060 was open with no notable canopy cover, so sunlight was accessible to the reach regardless of its orientation. The reach at GClOO runs east to west and was the only reach that was oriented in this manner except for sites BO040 and BO070. The east to west orientation of site GClOO coupled with relatively sparse canopy cover allowed aquatic vegetation to be exposed to sunlight longer than the other study sites that were typically oriented north to south. Although the reach sampled at BO040 shared the same orientation characteristics as GClOO, the pool habitat of BO040 was heavily shaded by dense canopy cover, which likely inhibited growth of J. americana by limiting light. Site BO070 is also oriented east to west, and similar to site GClOO canopy cover was sparse. The orientation and sparse canopy cover of site BO070 likely contributed to the relatively large amount of total biomass associated with Cladophora (375 gW) that was second only to site GClOO (927 gW). At BO070, periphyton rather than macrophytes, represented the majority of the biomass (Table 4). In comparing between habitat types (riffle and pool), the majority of periphyton biomass was obtained from the riffle habitat for all sites except BO060 (Table 5). For sites BO040, DC060, and MC055, the riffle and pool habitats contributed nearly equal amounts of the periphyton biomass. The riffle and pool habitats contributed to nearly equal amounts of the biomass obtained from macrophyte species at sites BO060 and BO085, while the majority of macrophyte biomass for the remaining mainstem sites originated from the pool habitats with the exception of site BO020. Discussion A large difference occurred in the estimated trophic status classifications between sites depending on whether periphyton or phytoplankton was used as the indicator (Figure 3). For periphyton, only four sites (BO070, BO080, DC060, and GClOO) were classified as mesotrophic or eutrophic using either maximum or mean values. For phytoplankton, all sites but DC060, MC055 and MCFARLAND ET AL. 15 Table 5. Percentage of total biomass obtained from algal and macrophyte species collected within each habitat type. Site Periphytic Algae Macrophyte Riffle (% biomass) Pool (% biomass) Riffle (% biomass) Pool (% biomass) BO020 91 9 100 0 BO040 51 49 34 66 BO060 20 80 54 46 BO070 62 38 46 54 BO080 88 12 13 87 BO085 70 30 52 48 BO090 64 36 17 83 DC060 57 43 38 62 GClOO 87 13 39 61 MC055 57 43 31 69 NC060 74 26 20 80 NC060 were classified as mesotrophic. This apparent discrepancy may in part be explained by competition between algal types. Streams dominated by periphytic algae may have less phytoplankton due to competition for nutrients, light or other limiting factors and vice versa. Also, the establishment of peri¬ phytic algae is dependent on the substrate of the streambed providing adequate attachment sites. Another factor, particularly in comparing the mainstem sites, is the upstream to downstream transport of algae. While periphyton can detach and move down¬ stream, this occurs predominately as a result of scouring associated with elevated flows. Phytoplankton are free-floating in the water column and move with the transport of water. Sestonic CHLA con¬ centrations showed a general decreasing pattern from more upstream sites (BO020, BO040 and BO060) to more downstream sites (BO085 and BO090). For periphyton, some of the lowest bio¬ mass estimates were indicated at the most upstream site (BO020). Nutrient concentrations of total N and total P were also generally higher in the upstream portion of the river than in the more downstream portion (Table 6). Excessive aquatic plant growth is often related to excessive nutrients, but defining target concentra¬ tions of N and P to help control algal (or macrophyte) impairment is Mean Phytoplankton CHLA (pg/L) Maximum AFDW Periphyton (g/m^) Mean AFDW Periphyton (g/m^) 16 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 14 12 10 8 6 4 2 0 o o o o O o IT) o o lO o CD o 00 CD 00 CD CM in CD O T— o o O O o O O o O o O O O o o o o o O O Q O 00 GO CD m DQ z QQ 00 n n n □ □□ ,=1 ] Value . O-M Boundary - M-E Boundary Figure 3. Comparison of potential trophic boundaries for periphyton and phytoplankton for streams based on ERA (2000) and work by Biggs (2000). O-M indicates oligotrophic to mesotrophic boundary and M-E indicate mesotrophic to eutrophic MCFARLAND ET AL. 17 Table 6. Basic statistics for total N and total P from routine grab samples collected between March 1, 2004 and June 30, 2005. N equals the number of observations. Standard Constituent Site Mean Median Deviation Minimum Maximum Number (mg/L) (mg/L) (mg/L) (mg/L) (mg/L) of Obs. Total N BO020 1.68 1.61 0.63 0.61 3.90 32 BO040 3.74 3.47 1.07 2.14 6.15 35 BO060 1.43 1.38 0.69 0.39 2.75 27 BO070 1.31 1.21 0.76 0.08 3.08 35 BO080 1.22 1.05 0.64 0.40 2.91 27 BO085 0.96 0.82 0.52 0.10 2.22 27 BO090 0.84 0.78 0.39 0.20 2.07 34 DC060 0.94 0.62 0.92 0.26 4.68 27 GCIOO 2.87 2.23 2.14 0.48 7.53 27 MC055 0.60 0.54 0.26 0.23 1.19 27 NC060 0.71 0.64 0.20 0.31 1.10 35 Total P BO020 0.40 0.41 0.18 0.09 0.85 32 BO040 1.52 1.11 0.91 0.56 3.66 35 BO060 0.44 0.42 0.21 0.01 1.06 27 BO070 0.21 0.21 0.11 0.06 0.49 35 BO080 0.13 0.10 0.10 0.01 0.40 27 BO085 0.10 0.08 0.09 0.01 0.38 27 BO090 0.08 0.07 0.07 0.01 0.32 34 DC060 0.10 0.06 0.11 0.02 0.44 27 GCIOO 0.11 0.06 0.14 0.01 0.60 27 MC055 0.06 0.06 0.03 0.01 0.15 27 NC060 0.06 0.06 0.03 0.01 0.13 35 difficult (TNRCC 2001). Chlorophyll-a concentrations of phytoplankton in the water column generally show a relatively close relationship with nutrient concentrations in the water column, particularly in lake systems (e.g., Carlson 1977; Vollenweider et al. 1980). For periphytic algae and macrophytes, a correlation with instream nutrient concentrations generally does not occur (Dodds et al. 2002). Macrophytes, in particular, obtain most of their nutrients from bank or bed sediments rather than the water column. The growth of maerophytes and periphyton may also be impacted by light availability, as noted earlier with regard to the abundance of J. americana at sites DC060 and GCIOO. The east to 18 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 west orientation of these two sites coupled with a sparse canopy cover is thought to have allowed more abundant macrophyte growth at these two sites than at other sites monitored in the watershed. While almost all macrophytes found were emergent or floating, turbidity within the water column can also limit light penetration limiting the growth of submerged macrophytes or periphyton. Although these stream sites can become quite turbid during storm events with total suspended solids (TSS) concentrations of several hundred mg/L, routine grab samples collected on a biweekly basis between March 2004 and June 2006 indicated median concentrations of TSS ranging from <4 to 12 mg/L. Although light availability within the water column is an important factor for aquatic plant growth, variations in TSS were not related to periphyton biomass. In an attempt to relate instream nutrient to periphytic CHLA concentrations, Dodds et al. (2002; 2006) evaluated almost 300 sampling events using a literature data set representing more than 200 rivers in North America and New Zealand. Dodds et al. (2002; 2006) found very weak (r < 0.32) but significant (a = 0.05) correlations between mean periphytic CHLA, as a measure of algal biomass, and instream nutrient concentrations. In earlier work, Dodds et al. (1997) proposed a saturation effect of nutrients on periphytic algae or a lack of further growth response when nutrient concentrations are high. Using breakpoint regression to define when saturation occurred, Dodds et al. (2002; 2006) determined saturation breakpoints for maximum benthic algal biomass concentrations at about 0.06 mg/L total P and 0.60 mg/L total N. For the North Bosque River sites, median stream concentrations were generally at or well above these breakpoints (Table 6). Because total N and total P concentrations were generally above the breakpoints established by Dodds et al. (2002), nutrients do not appear to be the dominant factor limiting periphyton and macrophyte biomass in the North Bosque River under normal conditions, although at times nutrients may be a limiting factor. Minimum total P concentrations at all but three sites were well MCFARLAND ET AL. 19 below the breakpoint of 0.06 mg/L total P and only two sites indicated minimum total N concentrations greater than the total N breakpoint of 0.60 mg/L (Table 6). Earlier work in the North Bosque River focusing on nutrient limitation of periphytic algae using instream periphytometers indicates the potential for nitrogen limitation at some locations in the upper third of the watershed and phosphorus limitation at locations in the lower portion of the watershed (McFarland et al. 2004). In this work (McFarland et al. 2004), dissolved inorganic nitrogen (DIN) concentrations rather than total N concentrations appear to be more strongly related to growth potential when nitrogen limitation was indicated. When nitrogen limitation occurred, DIN concentrations were 0.04 mg/L or less, while total N concentrations averaged 0.45 mg/L with only one observation of 0.74 mg/L greater than the total N breakpoint of 0.60 mg/L determined by Dodds et al. (2006). Minimum DIN concentrations (data not shown) from grab samples between March 2004 and June 2005 for several of the North Bosque River sites were below 0.04 mg/L. Most likely periphytic algae were opportunistic to changing water quality conditions and responded to changes in nutrient concentrations when other factors, such as light, were not limiting. Besides nutrient and light limitation, periphyton are expected to respond to flow conditions. Low to moderate levels of flow help in supplying a continuous, although often varying, source of nutrients to locations along a stream. Losses of periphyton, and to a lesser degree macrophytes, may result from scouring associated with elevated flows from large rainfall runoff events. These scouring events can reduce stream periphytic algal biomass even when nutrient concentrations are high, causing a disconnect between nutrient concentrations and biomass levels. Although scouring is known to reduce periphytic algal biomass, macrophytes are generally more resistant to scouring events because their roots are embedded into the streambed substrate 20 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 unlike periphytic algae, which are attached to the substrate surface. For sites with continuous flow data, the abundance of macrophytes and periphyton from each sampling event was overlaid to see if an impact from scouring events could be visually observed. While it appeared that there was some impact due to elevated flows between sampling events, a clear pattern could not be construed. The fairly long time periods between sampling events may have allowed regrowth to occur after a potential scouring event prior to the next sampling. Also, macrophyte and periphytic algal biomass indicated some seasonality with the lowest biomass levels most often occurring during the January 2005 sampling events. For periphyton, there was often a noted decrease in the summer months that is probably temperature related. Cladophora often dies off in midsummer when water temperatures increase above about 25°C (Dodds & Gudder 1992). Summary and Conclusions A survey of macrophytes and periphyton biomass at 1 1 locations within the North Bosque River system indicated a wide range of spatial variation over the two-year study. Instream nutrient concentrations did not appear to be the primary factor influencing the abundance of periphytic algae and macrophytes, because saturated rather than limiting nutrient conditions generally occurred along the river for both N and P. Other factors, such as light limitation, scouring, and seasonality, were likely more influential than nutrient concentrations in determining the biomass measured at any specific point in time. Trophic status as determined by measurements of phytoplankton differed greatly from trophic status as determined using measurements of periphyton indicating a need to evaluate eutrophication from more than one perspective. Marcrophytes and periphytic algae represent important and dynamic sinks as well as sources of nutrients within the North Bosque River system that need to be considered in evaluating water quality problems associated with aesthetics and eutrophication. Such efforts are taking place as indicated by EPA proposed stream nutrient criteria (EPA 2000), but more work is needed. MCFARLAND ET AL. 21 Acknowledgements Funding for this study was provided through the Texas Commission on Environmental Quality, Work Order 582-6-70859- 04 as part of the project, Monitoring to Support North Bosque River Model Refinement. We would like to thank Dr. Allan Nelson, associate professor in Biological Sciences at Tarleton State University, for his assistance in verifying the identification of some specimens and for allowing access to the Tarleton State University herbarium. Literature Cited Allan, J. D. 1995. Stream ecology: Structure and function of running waters. Kluwer Academic Publishers, Boston, MA, 388 pp. APHA. 1992. Standard Methods for the Examination of Water and Wastewater. 18‘^ edition. American Public Health Association, Washington, D.C., 1268 pp. Biggs, B. J. 2000. New Zealand Periphyton Guideline: Detecting, Monitoring and Managing Enrichment of Streams. Ministry for the Environment, New Zealand, Ref.ME359, 124 pp. Carlson, R. E. 1977. A trophic state index for lakes. Limnol. Oceanog., 22(2):361- 369. Chambers, P. A., R. E. DeWreede, E. A. Irlandi & H. Van DeMeulen. 1999. Management issues in aquatic macrophyte ecology: A Canadian perspective. Can. J. Bot., 77(4):47 1-487. Diggs, Jr., G. M., B. L. Lipscomb & R. J. O’Kennon. 1999. Shinners and Mahler’s illustrated flora of north central Texas. Center for Environmental Studies and Dept, of Biology, Austin College, Sherman, Texas, 1640 pp. Dodds, W. K. & D. A. Gudder. 1992. The ecology of Chladophora. J. PhycoL, 28(4):415-427. Dodds, W. K., V. H. Smith, & K. Lohman. 2006. Erratum: Nitrogen and phosphorus relationships to benthic algal biomass in temperate streams. Can. J. Fish. Aquat. ScL, 63(5):1 190-1 191. Dodds, W. K., V. H. Smith, & K. Lohman. 2002. Nitrogen and phosphorus relationships to benthic algal biomass in temperate streams. Can. J. Fish. Aquat. Sci., 59(5):865-874. Dodds, W. K., V. H. Smith, & B. Zander. 1997. Developing nutrient targets to control benthic chlorophyll levels in streams: a case study of the Clark Fork River. Water Res., 31(7):1738-1750. EPA. 2000. Nutrient Criteria Technical Guidance Manual: Rivers and Streams. Office of Water and Office of Science and Technology, Washington, D.C., EPA- 822-B000-002, 240 pp. Fritz, K. M., M. A. Evans, & J. W. Feminella. 2004. Factors affecting biomass 22 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 allocation in the riverine macrophyte Justicia americana. Aquat. Bot., 78(3):279- 288. Gibson, C. E. 1997. The dynamics of phosphorus in freshwater and marine environments. Pp. 119-135, in: H. Tunney, O.T. Carton, P.C. Brookes, and A.E. Johnston (eds.). Phosphorus Loss from Soil to Water. CAB International, New York, NY, 480 pp. Howell, H. H. 1975. Some ecological factors affecting the occurrence of water willow Justicia americana in Jessamine Creek, Kentucky. Trans. Ky. Acad. Sci., 36(l):43-50. Kiesling, R. L., A. M. S. McFarland, & L. M. Hauck. 2001. Nutrient targets for Lake Waco and North Bosque River: Developing ecosystem restoration criteria. Texas Institute for Applied Environmental Researeh, Tarleton State University, Stephenville, TEXAS, TR0107, 51 pp. McFarland, A., R. Kiesling, & J. Back. 2004. Using periphytometers to evaluate impairment due to nutrient enrichment. Texas Institute for Applied Environmental Research, Tarleton State University, Stephenville, Texas, TR0404, 51 pp. Newton, B. J. & W. M. Jarrell. 1999. A procedure to estimate the response of aquatic systems to changes in phosphorus and nitrogen inputs. National Water and Climate Center, United States Department of Agriculture, Natural Resources Conservation Service, Washington, D.C., 37 pp. Prescott, G. W. 1954. How to know the fresh-water algae. Wm. C. Brown Company, Dubuque, Iowa, 211 pp. SAS. 2000. The SAS system for windows. Ver. 8. SAS Institute, Cary, NC. Sosiak, A. 2002. Long-term response of periphtyon and macrophytes to reduced municipal nutrient loading to the Bow River (Alberta, Canada). Can. J. Fish. Aquat. Sci., 59(6):987-1001. TCEQ. 2003. Guidance for assessing Texas surface and finished drinking water quality data, 2004. Texas Commission on Environmental Quality, Surface Water Quality Monitoring Program, Austin, Texas, 93 pp. TNRCC. 1998. 1998 Clean Water Act Section 303(d) list and schedule for development of total maximum daily loads (TMDLs). Water Quality Division, Texas Natural Resource Conservation Commission, Austin, Texas, SFR-58. TNRCC. 2001. Two maximum daily loads for phosphorus in the North Bosque River for segments 1226 and 1255. Strategic Assessment Division, TMDL Team, Texas Natural Resouree Conservation Commission, Austin, Texas, 29 pp. Vollenweider, R. A., W. Rast, & J. Kerekes. 1980. The phosphorus loading concept and Great Lake eutrophication. Pp. 207-234, in: R.C. Loehr, C.S. Martin, and W. Rast (eds.). Phosphorus Management Strategies for Lakes. Ann Arbor Science Publications, Ann Arbor, Michigan, 490 pp. Welch, E. B., J. M. Jacoby, R. R. Homer, & M. R. Seeley. 1988. Nuisance biomass levels of periphytic algae in streams. Hydrobiologia, 157(2): 161 -168. MSM at: mcfarla(gtiaer.tarleton.edu TEXAS J. OF SCI. 60(l):23-32 FEBRUARY, 2008 DISTRIBUTION OF THE BANTAM SUNFISH, LEPOMIS SYMMETRICUS (PERCIFORMES: CENTRARCHIDAE), IN ARKANSAS Chris T. McAllister, Renn Tumlison and Henry W. Robison Department of Physical & Life Sciences, Chadron State College Chadron, Nebraska 69337, Department of Biology, Henderson State University Arkadelphia, Arkansas 71999 and Department of Biology, Southern Arkansas University Magnolia, Arkansas 71754 Abstract. -This study reports 77 new geographic and 15 new county records for the bantam sunfish, Lepomis symmetricus, collected between May 1988 and June 2007 in 30 counties of Arkansas. Detailed distributional data are included as well as an updated map showing the overall distribution of this fish in the state. Bantam sunfish were collected from all major drainages within the Gulf Coastal Plain and Mississippi Alluvial Plain physiographic provinces of Arkansas, including the Arkansas, Ouachita, Mississippi, Red, Saline, St. Francis and White rivers. Based on these new collections and historical records of the past, it is believed that populations of L. symmetricus in Arkansas are stable and do not warrant a protected legal status. The bantam sunfish, Lepomis symmetricus Forbes is a very small (< 9.0 cm) nongame centrarchid that ranges in the former Mississippi Embayment from southern Illinois to the Gulf Coast, including Gulf Slope drainages from Bay St. Louis, Mississippi, west to the Colorado River, Texas, and McCurtain County, Oklahoma (Smith 1979; Burr 1980; Page & Burr 1991; Miller & Robison 2004). The species inhabits shallow waters of lowland swamps, oxbow lakes and sluggish backwaters of creeks and bayous almost always in association with aquatic vegetation, submerged logs, and stumps over a detritus and muddy bottom (Burr 1977; Robison & Buchanan 1988). In Arkansas, this fish is reported to be more common in the southern part of the state than in the east, restricted to the Coastal Plain lowlands of all major drainages (Robison & Buchanan 1988). Black (1940) recorded the first occurrence of L. symmetricus in Arkansas based on collections from Monroe County (Maple 24 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Slough, 3.2 km east of Brinkley) and St. Franeis County (relief off the L’Anguille River, 1.6 km northeast of Palestine). Collections including bantam sunfish in Arkansas were so few that Buchanan (1974) treated the species as having an “undetermined status,” although he suggested that the species was “possibly endangered in Arkansas.” Since then, Robison (1975) added three more records including Big Comie Creek in Columbia County, a site in the Saline River drainage of Bradley County, and another from the Ouachita River backwater in Calhoun County. Loe (1983) provided two records of L. symmetricus from Nevada County. Robison & Buchanan (1988) provided 26 collection sites in all major drainages in Arkansas for the period 1960 to 1987, and two additional pre-1960 sites. Additional records for L. symmetricus were provided by Buchanan et al. (2003) who reported the species was uncommon in the Red River (upstream and downstream from the mouth of the Little River) of extreme southwestern Arkansas. The most recent records are those of Buchanan (2005) who used rotenone sampling to collect 180 bantam sunfish from six Arkansas reservoirs, including Champagnolle Creek, Dierks, Erling, Merrisach, Millwood, and Pool 2 of the Arkansas River. Methods and Materials To update the status of this uncommon sunfish this study sampled streams throughout Arkansas and mapped museum collection data. Between May 1988 and June 2007 bantam sunfish were collected with standard nylon seines (1.8 by 0.5 m and 2.7 by 0.5 m of 3.2 mm mesh) or dipnets. Fishes were preserved in 10% formalin and later transferred to 45% isopropanol. Specimens were field identified, verified in the laboratory, and vouchers deposited in the collections at Southern Arkansas University, Magnolia, Arkansas (SAU), the University of Louisiana-Monroe Museum of Natural History, Monroe, Louisiana (NLU), Henderson State University (HSU), the North Carolina State Museum of Natural Sciences, Raleigh, North Carolina (NCSM), and the University of MCALLISTER, TUMLISON & ROBISON 25 Arkansas at Fort Smith, Fort Smith, Arkansas (UA-FS). Detailed data provided on the 77 new sites are as follows: (county, specific locality [township, section, and range when available], date, museum accession number (if known), and number of specimens in parentheses). Material examined -The following is a listing of collection localities of L. symmetricus in Arkansas. County names followed by an asterisk (*) represent new county records. ARKANSAS COUNTY {n = 14): Cox Cypress Lake, 16.1 km NofBayouMeto(Sec. 2, T5S, R6W). 4 May 2001. NLU 77315 (2); Arkansas River, Merrisach Lake Park boat dock ramp. River Mile 14.47 (Sec. 6, T8S, R2W). 22 June 2004. NLU 78535 (4); Unnamed creek, 3.2 km N of St. Hwy. 165 (Sec. 25/26, T4S, R3W). 9 October 1996. NLU 74717 (3); Little LaGrue Bayou, 6.4 km NE of Almyra, upstream of weir (Sec. 12, T3S, R4W). 5 September 1996. NLU 74659, 74817 (8). ASHLEY COUNTY {n = 27): Hanks Creek, 0.3 km W of St. Hwy. 81 on U.S. Hwy. 82 (Sec. 28, T18S, R7W). 24 March 1989 and 9 May 1991. NLU 62310, 62315, 65393 (15); Hanks Creek at U.S. Hwy. 82W, 3.2 km S of Hamburg (Sec. 28, T18S, R7W). 29 April 1990. NLU 64413 (2); Hank’s Creek, 1.3 km W of St. Hwy. 81 on U.S. Hwy. 82. 15 June 1993. SAU (4); Thompson Creek, 11.3 kmNW of Crossett (Sec. 11, T17S, R9W). 5 July 1996. HSU 1343 (3); Bearhouse Creek at gravel road NE of Snyder (Sec. 22, T16S, R5W). 21 April 2000. SAU (1). Unnamed tributary to Chemin-A-Haut Creek at St. Hwy. 52 (Sec. 19, T18S, R7W). 21 April 2000. SAU (2 juveniles). BRADLEY COUNTY (n = 3): Moro Creek at Moro Bay State Park (Sec. 21, T16S, R12W). 10 June 2005. SAU (1); L’Aigle Creek at county road, 14.5 km S Hermitage (Sec. 18, T16S, RlOW). 10 June 2005. SAU (2). 26 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 CALHOUN COUNTY (« = 10): Locust Bayou at St. Hwy. 278, 14.5 km E of Camden (Sec. 30, TBS, R15W). 2 May 1990. SAU (1); Locust Bayou at St. Hwy. 278, 14.5 km E of Camden (Sec. 30, TBS, R15W). 1 June 2003. NCSM 36205 (1); Moro Creek at St. Hwy. 160 (Sec. 9, T16S, R12W). 18 June 2007. SAU (5); Champagnolle Creek at co. road (Sec. 27, TBS, R14W). 18 June 2007. SAU (3). CLARK COUNTY* {n = 8); Tupelo Creek at St. Hwy. 51, 3.2 km E of Arkadelphia. 13 March 1996. UA-FS (6); McNeeley Creek, 6.4 km S Beime off St. Hwy. 51 (Sec. 31, TIOS, R20W). 20 April 1997. HSU 2154 (1); Gurdon Lake. 17 August 2000. UA-FS (1). CLEVELAND COUNTY (n = 2): Saline River at TAR (Sec. 17, T1 IS, R9W). 7 July 1991. SAU (1); Big Creek at St. Hwy. 1 14 (Sec. 10, T9S, RlOW). 2 May 1996. SAU (1). CHICOT COUNTY* (n = 2): Overflow area E of levee on Mississippi River at end of St. Hwy. 144 (Sec. 21, TBS, RIE). 18 June 2002. SAU (2). COLUMBIA COUNTY (n = 9); Dorcheat Bayou at county road, 4.8 km SW of Philadelphia (Sec. 16, T18S, R22W). 16 September 1989. SAU (7 juveniles); Dorcheat Bayou at St. Hwy. 160, 6.4 km E of Taylor (Sec. 9, TBS, R22W). 19 May 2004. SAU (1); Horsehead Creek at U.S. Hwy. 79, 9.7 km E of Magnolia (Sec. 17,T18S,R20W). 14 May 2005. SAU(l). DALLAS COUNTY* (« = 7): East Tulip Creek at St. Hwy. 8 at Princeton (Sec. 34, T8S, RBW). 13 March 1996. SAU (5); Unnamed tributary to Freeo Creek at Ramsey (Sec. 5, TIOS, R14W). 29 April 2003. SAU (2). DESHA COUNTY* (n = 1); Canal 19 at St. Hwy. 138, E of Winchester (Sec. 6, T1 IS, R3W). 27 August 2003. NLU 78310 (1). MCALLISTER, TUMLISON & ROBISON 27 DREW COUNTY (n = 5): Cut-Off Creek at St. Hwy. 35, 1.1 km E of Collins (Sec. 31, T13S, R4W). 8 April 1994. SAU (2); Bayou Bartholomew at St. Hwy. 35, 6.4 km E Collins (Sec. 28, T13S, R4W). 28 September 1999. SAU (3). HEMPSTEAD COUNTY (n = 5): Millwood Lake at Saratoga Landing, 0.8 km W of Saratoga (Sec. 6, T12S, R27W). 10 August 1990. SAU (5). HOT SPRING COUNTY (« = 1): Saline Bayou, 4.0 km S Friendship (Sec. 23, T6S, R19W). 14 February 1997. HSU 2154 (1). HOWARD COUNTY* (n = 3): Saline River at county road, 3.2 km W of Schaal (Sec. 10, Til S, R28W). 12 June 2000. SAU (1); Mine Creek at county road, 1.6 km W of Toilette (Sec. 6, TllS, R29W). 1 1 August 1990. SAU (2). JEFFERSON COUNTY (n = 3): Yellow Lake on Pine Bluff Arsenal (Sec. 2, T5S, RlOW). 1 July 1999. SAU (2); Bayou Meto WMA, Wrape Plantation, roadside pool (Sec. 6, T6S, R5W). 8 May 2002. NLU 77306(1). LAFAYETTE COUNTY (n = 8): Bodcau Creek at county road, 1.6 km N Lewisville (Sec. 7, T15S, R23W). 16 March 1990 and 5 July 1992. SAU (2); Lake Erling at St. Hwy. 160 (Sec. 35, T19S, R23W). 24 September 1991. SAU (6). LINCOLN COUNTY* (n = 65): Bayou Bartholomew at Garrett Bridge (Sec. 6, TIOS, R5W). 3 June 1989. SAU (3); Oakwood Bayou at St. Hwy. 212, 8.0 km E of government road (Sec. 7, T9S, R7W). 17 April 2003. NLU 77785 (60); Bayou Bartholomew, 6.4 km N of Star City (Sec. 21, T8S, R7W). 7 July 2003. SAU (1); Bayou Bartholomew at St. Hwy. 293 (Sec. 15, T9S,R6W). 7 July 2003. SAU(l). 28 THE TEXAS JOURNAL OF SCIENCE- VOL. 60, NO. 1, 2008 LITTLE RIVER COUNTY (« = 13): Little River Relief at St. Hwy. 41, 6.6 km SW of Horatio (Sec. 10, TIOS, R32W). 25 May 1988. NCSM 37422 (5); Cypress Creek at St. Hwy. 234 in Winthrop (Sec. 7, TllS, R31W). 6 June 1989. SAU (1 juvenile); Flat Creek at St. Hwy. 234, 3.2 km E of Winthrop (Sec. 21, TllS, R31W). 10 July 1990. SAU (1 juvenile); Caney Creek at St. Hwy. 41, 4.0 km SW of Billingsley’s Comer (Sec. 28, TIOS, R32W). 1 March 1992. SAU (2); Little River backwater at U.S. Hwy. 71, 3.2 km N of Wilton (Sec. 24, TllS, R29W). 5 October 2001. SAU (4). LONOKE COUNTY* (n = 56): Indian Bayou at Tomberlin (Secs. 21/28, T2S, R8W). 14 July 2000. NLU 76916 (55); Arkansas River at River Mile 104, Willow Bar Cutoff off U. S. Hwy. 165, 0.4 km NW of Pilgrims Rest Church (Sec. 21, T2S, R9W). 13 July 2000. NLU 76772(1). MILLER COUNTY {n = 200): Mercer Bayou ramp off County Road 109, above lower stmcture (Sec. 19, T18S, R27W). 25 May 1988. NLU 75772 (3); Kelly Bayou at county road, 6.4 km SW of Doddridge (Sec. 9, T20S, R27W). 3 September 1990. SAU (1); West Fork Kelly Bayou at county road, 1.6 km S of Brightstar (Sec. 28, T18S, R28W). 3 September 1990. SAU (1). Mercer Bayou at gated structure, 7.6 km W of Beggy (Sec. 2, T18S, R28W). 24 May 1999. NLU 75765 (3); Upper Mercer Bayou, bendway between ramp and gate (Sec. 10, T18S, R28W). 25 May 1999. NLU 75785 (6); Middle Mercer Bayou, upper lateral tributary (Sec. 10, T18S, R28W). 25 May 1999. NLU 75791 (6); Lower reach of Mercer Bayou, ramp at County Road 109 (Sec. 30, T18S, R17W). 4 August 1999. NLU 75825 (4); Mercer Bayou, upper reach at gated stmcture, 7.6 km west of Beggy (Sec. 2, T18S, R28W). 1 1 August 1999. NLU 75811 (34); Mercer Bayou, middle reach at lateral trib. (Sec. 2, T18S, R28W). 1 1 August 1999. NLU 75820 (30); Mercer Bayou, upper reach at bendway between gate stmcture and ramp (Sec. 10,T18S,R28W). 1 1 August 1999. NLU 75815 (112). MCALLISTER, TUMLISON & ROBISON 29 MISSISSIPPI COUNTY* (« = 72): Mississippi River bar pit No. 17, 0.8 km E of Wilson (Sec. 22, T1 IN, RlOE). 4 August 1996 and 4 August 1997. NLU 74322, 74329 (10, 62). NEVADA COUNTY* (« = 27): Middle Creek, 14.5 km N of Prescott on St. Hwy. 19 (Sec. 27, T9S, R23W). 19 February 1983. NLU (1); Caney Creek, 4.8 km N of Bluff City on St. Hwy. 24 (Sec. 22, T1 IS, R20W). 19 February 1983. NLU (26). OUACHITA COUNTY (n = 5): En Core Fabre Bayou, 1.6 km N of Camden (Sec. 10, T13S, R17W). 22 August 1994. SAU (2); Bragg Lake, 2.0 km SE Bragg City at St. Hwy. 24 (Sec. 33, T12S, R18W). 12 April 1997. HSU 2055 (2); Freeo Creek at St. Hwy. 9, 8.9 km S of Dallas County line (Sec. 36, TllS, R16W). HSU 2184 (1). PHILLIPS COUNTY* {n = 8): Mississippi River borrow pits off St. Hwy. 20, 1 1.3 km S of Helena (Sec. 35, T3S, R4E). 21 July 1997. NLU 74145 (8). PIKE COUNTY* {n = 1): Unnamed slough, 1.6 km E jet. St. Hwys. 195 and 301 (Sec. 13, T9S, R24W). 6 April 1997. HSU 1557 (1). PRAIRIE COUNTY* (« = 3): Unnamed creek, 4.0 km W of Seidenstricker (Sec. 1 and 2, TIN, R6W). 4 September 1996. NLU 74630 (1); Hooks Canal at St. Hwy. 11, 4.8 km NE of Slovak (Sec. 20, TIN, R5W). 6 September 1996. NLU 74601 (1); South Fork, 2.4 km E of Hazen at U.S. Hwy. 70 (Sec. 21, T2N, R5W). 10 October 1996. NLU 74727 (1). PULASKI COUNTY (n = 3): Backwater area along St. Hwy. 365, 0.8 km N of Woodson (Sec. 8, T2S, RllW). 10 July 2005. SAU (3). 30 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 SEVIER COUNTY {n = 6): Rolling Fork River at St. Hwy. 24, 4.8 km W of Horatio (Sec. 28, T9S, R32W). 8 October 1989. SAU (3); Millwood Lake, Paraloma Landing at end of St. Hwy. 234 (Sec. 29, Ills, R28W). 18 June 2002. SAU (2); Little River at end of St. Hwy. 317, 4.8 km S of Brownstown (Sec. 25, TllS, R29W). 19 June 2002. SAU(l). UNION COUNTY* {n = 6): Bear Creek at St. Hwy. 160, 8.0 kmNE Mount Holly (Sec. 21, T16S, R17W). 17 March 1989. SAU (1); Tributary of Ward Creek at U.S. Hwy. 82, E of El Dorado (Sec. 15, T18S, R14W). 29 October 2000. NLU 77006 (2); unnamed tributary to Big Creek at U.S. Hwy. 82, 6.4 km W El Dorado (Sec. 30, T17S, R16W). 26 February 2002. SAU (1); Big Comie Creek at St. Hwy. 15 (Sec. 35, T19S, R18W). 3 March 2002. SAU (1); Lapoile Creek at U.S. Hwy. 82, 16.1 km NE Strong (Sec. 18, T18S, RlOW). 7 April 2003. SAU (1). WOODRUFF COUNTY* {n = 3): Bayou DeView at St. Hwy. 38, 1.6 km E of Cotton Plant (Sec. 33, T5N, R2W). 29 October 2001. NLU 78209 (3). Results And Discussion This study documents the collection of 576 (10 juvenile, 566 adult) L. symmetricus from 30 of 75 counties (40%) of Arkansas (Fig. 1). Of those, 176 specimens (31%) were taken on 11 August 1999 from Mercer Bayou in Miller County. The second largest collection, 62 L. symmetricus or 1 1 % of the total were collected on 4 August 1997 from a Mississippi River borrow pit in Mississippi County. Loe (1983), in his unpublished thesis, also collected 27 specimens of L. symmetricus from Nevada County not included in Robison and Buchanan (1988). Bantam sunfish were collected from all major drainages within the Gulf Coastal Plain (GCP) and Mississippi Alluvial Plain (MAP) physiographic provinces of Arkansas, including the Arkansas, Ouachita, Mississippi, Red, Saline, St. Francis, and White rivers. MCALLISTER, TUMLISON & ROBISON 31 Figure 1. Records of the bantam sunfish {L. symmetricus) in Arkansas. Open circles (o) are older records referenced herein; closed circles (•) are current records. Compared with the records in Robison & Buchanan (1988), this study adds 1 5 new county records of L. symmetricus, including the northeastemmost, southeastemmost, and southwestemmost range extensions in the state for Mississippi, Chicot, and Little River counties, respectively. Further, numerous other collections of the species are noted from new localities in counties with historical records. All specimens originated from habitats within the GCP and MAP, although several collections were taken near the interface of the GCP and the Ouachita Mountains uplift. Because collections presently indicate that L. symmetricus is more widespread and common than was previously known, the authors believe that its populations in Arkansas are stable and do not warrant a protected legal status. Acknowledgments Special thanks are extended to Drs. T. M. Buchanan (UA-FS), N. H. Douglas (ULM) and W. C. Starnes (NCSM) for use of their previously unpublished Arkansas records of Lepomis symmetricus. 32 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Appreciation is extended to numerous collectors, especially T. M. Buchanan, N. H. Douglas and W. C. Starnes. Also, thanks to previous SAU Vertebrate Natural History classes, and former SAU students Jan Rader, Christa Brummett, Nick Covington, and Ken Ball, and HSU ichthyology classes, and former HSU students Kevin Funderbork and Jeannie Thompson for assistance in collecting. We also thank the Arkansas Game and Fish Commission for providing scientific collecting permits to HWR and RT. Literature Cited Black, J. D. 1940. The distribution of the fishes of Arkansas. Unpublished Ph.D. dissertation, Univ. Michigan, Ann Arbor, 243 pp. Buchanan, T. M. 1974. Threatened native fishes of Arkansas. Pp. 67-92, in Arkansas Natural Area Plan (T. Foti, Ed.), Arkansas Department of Planning, Little Rock, 248 pp. Buchanan, T. M. 2005. Small fish species of Arkansas reservoirs. J. Arkansas Acad. Sci., 59:26-42. Buchanan, T. M., D. Wilson, L. G. Claybrook & W. G. Layher. 2003. Fishes of the Red River in Arkansas. J. Arkansas Acad. Sci., 57: 18-26. Burr, B. M. 1977. The bantam sunfish, Lepomis symmetricus: Systematics and distribution, and life history in Wolf Lake, Illinois. Illinois Nat. Hist. Surv. Bull., 31:437-466. Burr, B. M. 1980. Lepomis symmetricus. P. 603, in Atlas of North American freshwater fishes (D. S. Lee, C. R. Gilbert, C. H. Hocutt, R. E. Jenkins, D. E. McAllister & J. R. Stauffer Jr., eds). North Carolina State Museum of Natural History, Raleigh, 854 pp. Loe, R. A. 1983. A subsequent study of the fishes of the Little Missouri River, west central Arkansas. Unpublished M.S. Thesis, Northeast Louisiana Univ., Monroe, 139 pp. Miller, R. J. & H. W. Robison. 2004. Fishes of Oklahoma, 2"^ Ed. University of Oklahoma Press, Norman, Oklahoma, 450 pp. Page, L. M. & B. M. Burr. 1991. A field guide to freshwater fishes. North America north of Mexico. Houghton Mifflin Co., Boston, Massachusetts, 432pp. Robison, H. W. 1975. New distributional records of fishes from the lower Ouachita River system in Arkansas. Proc. Arkansas Acad. Sci., 29:54-56. Robison, H. W. & T. M. Buchanan. 1988. Fishes of Arkansas. Univ. Arkansas Press, Fayetteville, 536 pp. Smith, P. W. 1979. The fishes of Illinois. Univ. Illinois Press, Urbana, 314 pp. CTM at: drctmcallister@aol.com TEXAS J. OF SCI. 60(l):33-44 FEBRUARY, 2008 A POSITIVE CONTROL FOR DETECTING HETERODUPLEXES IN DGGE FOR MICROBIAL COMMUNITY FINGERPRINTING Andrew Syvyk, Armen Nalian, Michael Hume* and Alexandra Martynova-VanKley Department of Biotechnology, Stephen F. Austin State University Nacogdoches, Texas 75965 and *USDA, ARS, SPARC, Food and Feed Safety Research Unit College Station, Texas 77845 Abstract.-Denaturing gradient gel eleetrophoresis in combination with PCR has found a wide application for analysis of genetic variants in many disciplines of bioscience, especially evaluation of microbial diversity in environmental samples. However, amplification of an environmental sample containing multiple DNA species can lead to formation of heteroduplexes (HDs), a PCR artifact. Appearance of HDs on the denaturing gradient gel can lead to a skewed interpretation of results and inaccurate conclusions. The present study is designed to develop a simple methodology to verify presence or absence of heteroduplexes in PCR samples which in conjunction with ‘reconditioned’ PCR can be used as an effective control to indicate presence and positions of HDs during DGGE analysis. Identification of HDs can allow effective design of reconditioned PCR by varying template and primer amounts and PCR cycle number. Denaturing gradient gel electrophoresis (DGGE) is a powerful technique for separation of double-stranded DNA molecules of nearly identical size according to their nucleotide composition. This technique has a wide range of applications, from detection of point mutations to evaluation of microbial community diversity. DGGE exploits the phenomenon of discrete melting behavior of double stranded DNA. PCR-amplified fragments are separated during electrophoresis according to their melting behavior on a polyacrylamide gel with a linearly increasing denaturant gradient. The resulting DGGE band patterns reflect sequence diversity in a given sample. However, during the final PCR cycles when the concentration of primers is low, heteroduplexes (HDs) can form. HDs result in “phantom bands” on the denaturing gradient gel and often are erroneously interpreted as additional DNA species or sequence variants (Acinas et al. 2005). This can lead to overestimation of sequence population diversity. Cloning of PCR products from a mixed DNA template sample can exacerbate the 34 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 problem. The non-directed mismatch repair system of E. coli generates a number of novel sequences from a single HD clone (Ruano 1992; Qiu et al. 2001). Other screening methodologies which utilize PCR products such as RFLP and Thermal Gradient Gel Electrophoresis (TGGE) are prone to the same problem (Osborn & Moore 2000). In the past there were a number of reports of PCR artifacts pertaining to DGGE. With the use of constant denaturing capillary electrophoresis, formation of HDs was demonstrated and a simple "reconditioning PCR" suggested (Jensen 1993; Thompson & Marcelino 2002) to reduce the number of HDs produced. In another study to eliminate HDs, samples were treated by T7 endonuclease (Qiu et al. 2001). Surprisingly, a recent literature search revealed neither evidence of research being conducted according to suggested improvements nor a general awareness of the problem within the DGGE community. This is probably due to a fact that in spite of recommended methods, researchers lack control measures to judge the absence or presence of HDs. Therefore, this study attempts to identify the presence and positions of HDs on DGGE gels and suggest a simple methodology which can serve as a positive control to indicate the presence of HDs on DGGE gels. Material and Methods DNA extraction and amplification -ThvQQ groups of samples were prepared from 119bp fragments of 18S rDNA from the protistian parasites Eimeria tenella (A), Eimeria maxima (B) and Eimeria acervulina (C). Group I contained seven mixes: A only, B only, C only, AB, AC, BC and ABC. Group II contained group I mixes which were subjected to heating to 92°C and then allowed to cool to room temperature. The heating treatment was done in order to mimic one denaturing step of the PCR cycle in absence of primers. Group III was prepared by PCR (50 pi final volume) using group I mixes as templates (0.5 pi) with Eimeria species- specific primers (forward primer containing GC-clamp 5’- SYVYK ET AL. 35 GCCCGCCGCGCCCGCGCCCGTCCGCCGCCCCCGCCCGGATT AGATACAAAACCAACCC-3’, and reverse primer 5’-GCTGATA GGTCAGAAACTTG-3’, 0.8 ng/pl final concentration for both primers). The amplification process was carried out using 25 pi of JumpStart™ REDTaq® ReadyMix™ PCR Reaction Mix (Sigma Chemical Co., St. Louis, MO) in a Mastercycler (Eppendorf, Scientific Inc., Westbury, NY) according to the following program: initial denaturation at 94°C for 3 min followed by 35 cycles of denaturation at 92°C for 45 s, annealing at 60°C for 35 s, and an extension at 72°C for 2 min. The final extension was performed at 72°C for 7 min. To demonstrate formation of HDs as a function of PCR cycles, a sample containing all three fragments (ABC), was amplified using the same set of primers and amplification conditions as described above but with a reduced number of cycles (20, 16, 15, 14, 13 and 12 cycles). All PCR products were purified with a Wizard PCR purification kit (Promega, Inc., Madison, WI) and run on a 1% agarose gel to check for quality of amplification. DNA extraction and amplification from a field sample -¥qc^\ field samples from a poultry farm located in east Texas were collected and used for DNA extraction. The birds were infected with several species of Eimeria. The extractions were carried out using the QIAamp DNA Stool Mini Kit QIAGEN (QIAGEN, Valencia, CA). The extracted DNA (0.5ul) was used as a template for PCR reactions with the same set of primers and PCR conditions as described above. The product of the PCR reaction (0.5ul) was used as a template to carry out three cycles of reconditioned PCR. One field sample was used to carry out a series of PCR reactions with an increasing number of cycles from 11 to 23. DGGE analysis -VolydiCYyXdixmdQ gels (8%, 0.75mm thick) were prepared using a 35-45% gradient of urea-formamide mix (Myers & Maniatis 1987). Electrophoresis was conducted in IxTAE buffer at 60°C using a DGGE-2001 apparatus (CBS Scientific Co., Del Mar, CA). After electrophoresis, the gels were stained with SYBR Green I and subsequently photographed using a BioRad Imager 36 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 System equipped with a Gel Doc XR camera and Quantity-One software (BioRad Inc., Hercules, CA). Results Formation of HDs.-Wq demonstrated the formation of HDs using a simple system on denaturing gradient (DG) gel. The model system contained the PCR products from the single A, B, and C fragments and their mixture ABC which we loaded on the DG gel (Group I) to show that no heteroduplexes were present in the original single template PCR products or in their mixtures. The heat-treated samples (Group II) along with the PCR products (Group III) were also loaded on the gel (Figure 1). As revealed by DGGE analysis, the PCR products from mixtures BC, AC, and AB contained in addition to the expected two original bands, two HD bands located higher on the gel than their parental bands (Group III, Figure I). In the same group, the PCR product of the ABC mixture template showed five bands in addition to the three parental bands (Six HD bands were expected, however due to co-migration only five bands were visible). The HD bands represent HDs derived from all possible matches of the parental single strands. By matching the band patterns from the dual mixture samples (Group III) the origin of all the HD bands can be traced back to their corresponding parental bands. Group II samples which included the heat treated mixtures (ABC, BC, AC, and AB) showed striking similarity to their mixed template PCR counterparts from Group III. Moreover, by comparing bands from Group I and Group II or III it is possible to clearly identify HDs by a simple elimination of common bands. Elimination of /TO^.-The presence of identical HDs in Group II and Group III as shown on Figure I leads to a conclusion that a single denaturation step in absence of primers is sufficient for detection of HDs. After the denaturation step during the last cycles of PCR when the primer: template ratio is low, single strands can rehybridize and form all possible combinations of hetero- and homoduplexes (Ruano 1992; Jensen 1993). To determine the first SYVYK ET AL. Group II Group III 37 (O TD C CO m E o +-» c CO -C a. Group 1 AA AA AA A A B B B BBB BB G C CCC CCC Figure 1. Formation of HD bands by PCR and heat treatment. Group I contained amplicons from single template PCR products and their mixture. Group II represents Group I samples which were subjected to heat treatment at 92°C and then cooled to room temperature. Group III samples contained PCR products obtained from mixed templates. Patterns of HD migration: a - co-migration (HDs migrate as a single band),, p - double-band migration (HDs migrate close to each other), y - separate migration (HDs migrate as two separate bands). PCR cycle when HDs ean be deteeted, PCR products of the ABC mixture obtained from six amplification reactions with decreasing number of cycles were analyzed by DGGE (Figure 2). Loading Set I contained equal volumes of PCR products from every reaction. Loading Set II contained adjusted volumes of the same set of PCR products as in Loading Set 1. The volumes were adjusted to emphasize that the disappearance of the HD bands was due to reduction of PCR cycles and not due to a lower amount of PCR product. Therefore, volumes of samples with less PCR product 38 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 35% CD o c 0 ID CD O D) C 13 •+-J CD C 0 Q 45%. Loading Set I Loading Set II Number of Cycles Number of Cycles 20 16 15 14 13 12 20 16 15 14 13 12 t; Figure 2. Elimination of HD bands by reduction of PCR cycles. PCR product from a mixture of ABC loaded on a DG gel in two groups. Group I contained equal volumes of all the PCR products (2.5 pi). To confirm the absence of HD bands in some samples, the volumes of samples in Group II were adjusted to maximize the volume of the loaded sample and to be inversely proportional to the PCR cycle number. were increased to confirm the absence of HDs. Detection of HDs at different PCR cycles is complicated by different parameters such as PCR efficiency and initial copy number of template DNA therefore the proposed control measure is required to judge the presence, absence and position of HDs. Environmental sample -E\\mmdi\Aori of HDs by reduction of PCR cycles was tested using a field sample of unknown template composition. A series of PCR reactions was performed with reduced numbers of cycles from 22 to 1 1 (Figure 3). Sixteen cycles SYVYK ET AL. 39 A NumberofCycles M 11 12 13 14 15 16 17 18 19 20 21 22 HT M 30 30 30 30 20 16 17 16 12 10 8 7 6.5 Amounts loaded on the gel[|iM Figure 3. Elimination of HD bands in a field sample by cycle reduction: (a) Two percent agarose gel of PCR products obtained with different cycle numbers using field sample DNA extract. The gel was stained in ethidium bromide. Each well contains 2 pi of PCR product, (b) Parallel denaturing gradient gel of the same samples as in Figure 3a. Volumes of samples resulting from fewer PCR cycles were increased to demonstrate absence of heteroduplexes. The last lane (HT) contained a heat-treated PCR product. Heteroduplexes are marked with arrows. were required to reaeh the deteetion limit on agarose gel (Figure 3, A). The DGGE analysis of the corresponding reactions demonstrated the gradual appearance of HDs in the upper part of the gel (Figure 3, B). No HD bands were detected before the 18th cycle. Thus, a controlled reduction in the number of PCR cycles 40 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 35% (ts (0 2 2 cu Qj a> a. *Q. Q. Q. Q. CL £ E £ E E £ ro (/» Ml . I ^ W-m , k-- m ra ■ P i f4 I i & 3 4 _ 45% Figure 4. Elimination of HD bands in field samples by reconditioned PCR: Parallel denaturing gradient gel was loaded with six heat treated samples and their respective PCR reconditioned samples. Reconditioned samples are marked with an asterisk. HDs can be identified by comparing the heat treated samples with the adjacent reconditioned sample lanes. Reconditioned PCR was performed by running another three cycles of PCR using pi of original PCR product as template. demonstrates formation of HDs as a function of PCR cycles. However, elimination of HDs by the PCR cycle reduction strategy is not practical for a large number of samples. Elimination of HDs by PCR reconditioning carried out with six field samples (Figure 4) showed a marked decrease in number of bands in reconditioned samples. Most of the HDs as expected were found in the upper part of the gel, however several were found in the lower part. In addition, we noted several bands in the lower part of the gel which were present in the reconditioned samples and absent in the heat treated samples pointing out that any given DNA species can disappear by forming HDs. SYVYK ET AL. 41 Discussion The ability of denatured nueleie acid molecules to renature or hybridize forms the basis for numerous methods and applications including southern blot, northern blot and PCR. This same property, if not accounted for, can lead to the misinterpretation of results (Thompson & Marcelino 2002). In the present study using DGGE, clear evidence was obtained of quantitative and qualitative parameters that effect HD formation. These are not the artifactual bands which can form during the late PCR cycles in a single template containing samples due to secondary structure formations (Janse et al. 2004). This study demonstrates by using a simple model system where mixtures of double stranded homologous molecules were subjected to heating and cooling, that any two homoduplexes can form heteroduplexes in a single step. For instance A-A^, where A” designates the antisense and A^ the sense strand of DNA respectively, and require a single step of denaturation- renaturation to produce two HDs A'B^ and A^B" (Group II, Figure 1). Quantitatively, the number of possible hybrids will depend on the number of original homoduplex molecules present in the mixture. The maximum number of possible HDs can be calculated according to the formula: Nht=Nhm(Nhm-l) where Nht is the number of HDs and Nhm is the number of homo-duplexes. Thus, two homoduplex species during rehybridization will generate two additional heteromolecules; three will produce six, four - twelve and so on. Qualitatively, the structure of these hybrids as compared to the structure of homoduplexes has shape-distorting mis- match(es); thus, they melt under milder denaturing conditions and have different rates of migration on the denaturing gradient gel. It is therefore safe to say that HD bands on the denaturing gradient gel will always occur above the parental homoduplex bands. However, in a mixed template sample it is possible to have some homoduplex bands with lower melting temperature than some of the HDs. So, bands appearing on the upper part of the gel cannot always be 42 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 discarded as HD bands. Therefore, to identify HDs on the gel use of a positive control is required. Stability of the new hybrid molecules depends on the GC content of parental sequences and on their similarity. HDs can exhibit three different patterns of migration (Figure 1): (1) Co¬ migration (HDs migrate as a single band), (2) Double-band migration (HDs migrate close to each other), (3) Separate migration (HDs migrate as two separate bands). Due to the unstable structure of HDs, some may denature without forming a distinct band or may disappear after some time during the electrophoresis and leave smears. However, the number and amount of HDs can be significant and, if co-migrating, they can provide a strong fluorescent signal on a gradient gel and lead to false conclusions regarding total number of sequences and DNA species diversity. Using the A, B and C fragments in all possible combinations as a template for four PCR reactions this study has demonstrated that HD formation occurs in absence of primers during a single denaturation step (Group II, Figure 1). During the first cycle of PCR, when different but closely related templates with identical priming sites are present, double the number of original homoduplexes is generated. As PCR pro¬ gresses, the number of primers decreases exponentially at the same rate as the new strands are synthesized. Therefore, each consecutive PCR cycle creates an equilibrium shift from primer- single strand complex formation towards the formation of homo- and heteroduplexes. At a low concentration of primers or in their absence, there is an equal probability of formation of homo or heteroduplexes upon renaturation. The amount of heteromolecules depends entirely on the amount of primers, which are better competitors for hybridization sites. The presence of a GC clamp on one or both primers is required for DGGE resolution and provides favorable conditions for HD formation. As demonstrated in Figure 1 (Group II), a single denaturation-renaturation step is sufficient to form HDs. Such a condition repeatedly occurs during late cycles of PCR and it results in an even redistribution of homo and HDs. SYVYK ET AL. 43 Formation of HDs in multi-template PCR is inevitable and it contributes to the ‘Cot effect’ allowing amplification of less dominant DNA species during late cycles of PCR (Mathieu-Daude etal. 1996). Several methods have been proposed to reduce or eliminate HDs to obtain a representative picture of DNA species diversity by DGGE analysis. Some of the unstable HDs could be eliminated by increasing denaturant concentration in the gel although this may not always remove all the hybrids (Qiu et al. 2001). Elimination of HDs by the T7 endonuclease I (Lowell 2000) demands strict experimental conditions with respect to enzyme concentration and incubation time which differ from sample to sample and can not be predicted (Qiu et al. 2001). PCR reconditioning (Thompson & Marcelino 2002) can be helpful, but with the limitation that a ten fold increase in primer concentration will only provide enough primers for three additional PCR cycles and a hundred-fold increase would allow for only five additional PCR rounds before depletion of primers. In addition, PCR amplification depends on many factors such as primer efficiency, concentration of inhibitors and initial concentration of the template. Therefore, conditions for reconditioning PCR must be determined empirically. This study proposes a simple positive control which can be used to control for the presence of HDs in a PCR sample containing multiple templates. Heating an aliquot of the individual PCR sample to 96°C followed by an on-bench cooling will generate all possible HDs. By comparing the DGGE band profiles of the original PCR product and the heat treated sample, a researcher can reach a grounded conclusion about the presence, absence and location of HD bands on the gel (Figure 4). Hopefully this finding will help the DGGE community to generate reliable data and make interpretation of DGGE results easier. Acknowledgments We would like to thank Dr. Christian Zwieb (University of Texas Health Science Center at Tyler, Texas) and anonymous 44 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 reviewers for critical review and comments on an earlier draft of this manuscript. This work was supported by Mr. Stan Decker and Stephen F. Austin State University ORSP grant. Literature Cited Acinas, S. G., R. Sarma-Rupavtarm, V. Klepac-Ceraj & M. F. Polz. 2005. PCR induced sequence artifacts and bias: insights from two 16S rRNA clone libraries constructed from the same sample. Appl. Environ. Microbiol., 71(12):8966- 8969. Janse L, J. Bok & G. Zwart. 2004. A simple remedy against artifactual double bands in denaturing gradient gel electrophoresis. J, Microbiol. Methods, 57(2):279-281. Jensen, M. A. & N. Straus. 1993. Effect of PCR conditions on the formation of heteroduplex and single-stranded DNA products in the amplification of bacterial ribosomal DNA spacer regions. PCR Methods Appl., 3:186-194. Lowell J. L. & D. A. Klein. 2000. Heteroduplex resolution using T7 endonuclease I in microbial community analyses. Biotechniques, 28:676-678 Mathieu-Daude F., J. Welsh, T. Vogt & M. McClelland. 1996. DNA rehybridization during PCR: the 'Cot effect' and its consequences. Nucleic Acids Res., 24(ll):2080-2086. Myers, R. M., T. Maniatis & L. S. Lerman. 1987. Detection and localization of single base changes by denaturing gradient gel electrophoresis. Meth. Enzymol., 155:501-527. Osborn, A. M., E. R. B. Moore & K. Timmis. 2000. An evaluation of terminal- restriction fragment length polymorphism (T-RFLP) analysis for the study of microbial community structure and dynamics. Environ. Microbiol., 2:39-50. Qiu, X. Y., L. Y. Wu, H. S. Huang, P. E. McDonel, A. V. Palumbo, J. M. Tiedje & J. Z. Zhou. 2001. Evaluation of PCR-generated chimeras: Mutations, and heteroduplexes with 16S rRNA gene-based cloning. Appl. Environ. Microbiol., 67:880-887. Ruano, G. & K. K. Kidd. 1992. Modeling of heteroduplex formation during PCR from mixtures of DNA templates. PCR Methods Appl., 2:1 12-1 16. Thompson, J., L. Marcelino & M. F. Polz. 2002. Heteroduplexes in mixed-template amplifications: formation, consequence and elimination by 'reconditioned PCR'. Nucleic Acids Res., 30(9): 2083-2088. AMVK at: avankley@sfasu.edu TEXAS J. OF SCL 60(l):45-56 FEBRUARY, 2008 SURVIVORSHIP OF RADIO-TRANSMITTERED URBAN WHITE- WINGED DOVES ON THE EDWARDS PLATEAU, TEXAS Michael F. Small, John T. Baccus, and T. Wayne Schwertner* Department of Biology, Wildlife Ecology Program Texas State University, San Marcos, Texas 78666 and *Texas Parks and Wildlife Department, 201 Westmoreland Mason, Texas 78656 Abstract -This study evaluates survivorship of White-winged Doves (Zenaida asiatica) with subcutaneously implanted radio-transmitters in Mason, Texas from January to March of 2006. Attempts were made to compare survivorship of transmittered doves (0.70 ± 0.135) with untransmittered doves using mark-recapture methodology in program MARK. No mark-recapture survivorship models had adequate fit for comparisons. Consequently, annual survivorship was estimated from period survivorship data for transmittered doves. This study concluded that the use of radio-transmitters had a detrimental effect on White-winged Dove survival for this particular study. George et al. (2000) reported highly variable survival rates for White-winged Doves {Zenaida asiatica) in Texas and New Mexico from 1960 to 1978 using banding data. However, no recent reliable estimates of survival exist for White-winged Doves, and no comparisons of survival estimation methods have been conducted for this species. Radio-telemetry has become a frequently used tool for studying doves in the U.S. in recent years (Schulz et al. 1998; 2001; Schaefer et al. 2004; Small et al. 2005; Berdeen & Otis 2006). Captive studies of both Mourning Doves {Zenaida macroura) and White¬ winged Doves indicated subcutaneous implantation of transmitters were the preferred method of attachment (Schulz et al. 2001; Small et al. 2004b). A study of free-ranging Mourning Doves in South Carolina indicated implanted radio-transmitters did not detri¬ mentally affect survival from hunter harvest (Berdeen & Otis 2006). However, doves in that study showed higher survival rates when held 24 h post-implantation prior to release. 46 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 This study estimates annual survival for free-ranging White¬ winged Doves with subcutaneously implanted radio-transmitters, and tests the concept put forth by Guthrie & Lusk (2004) that using radio-transmitters for estimating survivorship may negatively bias estimates. This study provides an evaluation of the efficacy of this transmitter attachment technique in wild White- winged Doves. All activities were conducted in accordance with Texas State University-San Marcos lACUC approval #06-05CC59736D, state permit #SPR-0890-234, and federal permit #06827. Methods Study area -This study was conducted in Mason, Texas in the Edwards Plateau ecoregion (Gould et al. 1960). Mason encompasses 958.3 ha with a population of about 2,211 (City- data.com 2005). Dominant tree species in Mason include oaks {Quercus sp.) and pecan {Carya illinoinensis) mixed with ornamentals and an understory of ornamental shrubs and turf grass. Mason is inhabited by a variety of mid-size predators including domestic cats {Felis domesticus), raccoons (Procyon lotor), opossum (Dasypus virginianus), domestic dogs {Canis familiaris), and hawks (Accipiter sp.). White-winged Doves in Mason occur in significantly higher densities in discreet core areas of town (Schwertner & Johnson 2006). Also, backyard bird feeders are abundant in Mason and are frequently used by White-winged Doves. Areas of White-winged Dove habitat in Mason were delineated using ArcGIS 9.1 (Environmental Systems Research Institute, Inc., Redlands, CA, USA). The 1992 National Land Cover Data (U.S. Geological Survey 1999) were used to identify areas of urban land cover. These areas were then buffered at 500 m and the resulting polygons defined as White-winged Dove habitat (Schwertner & Johnson 2006), thus designating the study area. SMALL, BACCUS & SCHWERTNER 47 Radio-transmittered doves -Yoviy-fom White-winged Doves were implanted with subcutaneous radio-transmitters (Advance Telemetry Systems, Isanti, Minnesota) between 13 January and 11 March 2006 following the field implantation procedure described by Small et al. (2004a). Transmitters were 41.83 (±0.05) by 26.83 (±0.05) by 7.62 (±0.05) mm with 15.24 (±0.01) cm antennas and had a mean weight of 3.41 g. {SE = 0.01) (<3.0% dove body weight). Transmitters had a range of 1.14 km (±0.02) and an expected battery life of 334 d. Doves were released upon regaining a lucid state. Transmittered doves were monitored 5 days/week from time of implantation until 15 May 2006 (123 days) over the entire study area. White-winged Doves in Mason are urban obligate nesters and roosters (Schwertner & Johnson 2006). Although searches for doves were conducted during all periods of the day, because the study area was small in relation to transmitter range 2-3 days/week included monitoring radio-transmitter signals in early morning before daylight when the doves were assumed to be stationary. This enabled full coverage of the study area and consequently the conclusion that an individual had left the study area when a signal was not received for >5 days. On these occasions it was not possible to determine where the individual had relocated (Small et al. 2006b). There was no reason to believe that any transmitters failed during the course of this study. In two previous radio¬ telemetry studies with monitoring lasting 120 days (Small et al. 2005) and 96 days (Small et al. 2006a), all transmitters that failed gave off a distinctive whine signal for several days pre-failure, which did not occur during this study. In this study, all transmitters from known predations were found because they continued to transmit a signal (including one transmitter that had passed through the digestive tract of a cat and was embedded in the scat). Consequently, there was no evidence that any transmitters were destroyed by predators. 48 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 In the event of mortality, eause of death was determined by observation of predators with the dove or by the eondition of the eareass at the transmitter reeovery site and the presenee of predator seat, fiir, or feathers. Estimated pooled survivorship for transmittered White-winged Doves was determined using Kaplan-Meier produet limit estimator (Kaplan & Meier 1958; Pollock et al. 1989; White & Garrott 1990). White-winged Doves were categorized as dead on the day of transmitter recovery. Because the fate of missing doves was unknown they were right censured on the last day a signal was received. The Kaplan-Meier method bases survivorship estimates on survival days only. Consequently, all individuals which die or leave the study (regardless of their fate) are treated the same and right-censored. Analyzing the data without including those individuals of unknown fate (i.e., those believed to have left the study area) would be a misuse of the technique and provide erroneous results. Survivorship estimates were obtained by extrapolating period survival to annual survival. While this requires the typically unreasonable assumption of equal survivorship across periods, there is sufficient natural history information available on White-winged Doves to validate the method (see discussion). Consequently, mean daily survival rate (5'^) was estimated by multiplying estimated survivorship for the period being monitored (Spr) by the number of days (/) in the sample period (in this case 123 days). Thus the mean daily survival estimate was: s,=(Sp,)(ty Annual survival rate (So) was then estimated using the formula ^,-(^^)/(365), the probability of surviving one year (365 days). SMALL, BACCUS & SCHWERTNER 49 These data were assessed for demographie plausibility by ealculating the demographic age ratio (R) for juvenile to adults required for population stability (finite growth multiplier = 1.0) (Guthrey 2002; Guthery & Rusk 2004). In other words, given the estimated annual survivorship, how many young would have to be produced in order to maintain a constant population size? This can be determined from the following formula, R = (l/Sayh where R refers to the number of young needed to be produced to maintain the population size, (1/iS'^) represents the proportion which survive one year. By subtracting the proportion expected to survive one year from 1.00, the number of young needed to be recruited into the population can be estimated. An R = 2 {2 young produced/adult) value was considered conservatively plausible and 2 < R < 4 was considered possible but unlikely for telemetry derived survival rates (Small et al. 2005; 2006b). Values > 4 were considered physiologically unrealistic. Mark-recapture -\n addition to monitoring transmittered White¬ winged Doves, untransmittered doves were trapped and banded and mark-recapture methodology employed to estimate survivorship of untransmittered doves to those with radio-transmitters. White¬ winged Doves were trapped using modified wire funnel traps (Reeves et al. 1968) on 78 days during the period in which transmittered doves were monitored. Time between the first trap and last trap days was 118 days. During this period 476 individual doves were captured and recaptured and each dove’s capture history recorded. Program MARK (White & Burnham 1999) was used to evaluate survivorship models. Models were constructed and run using designated trap occasions ranging from 1 to 24 days. Program MARK models indicated the most parsimonious models were obtained using a trap occasion consisting of 12 days. Thus, 123 50 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 days/12 days yielded 10.25 trap occasions. Pooled survivorship for untransmittered doves was estimated for the period 18 January through 15 May 2006 using recaptures only models with a sin link function and 2^^ part variance estimation in Program MARK. Four models were constructed using parameters for survival, O, and probability of capture, p, with and without time dependency. Akaike Information Criterion corrected for small sample size (AICc) was used to select the most parsimonious model, 0(.) p{t) (Burnham & Anderson 2002). Parameter index matrices (PIMs) of the selected model were adjusted and all values in the O PIM set to 1 and values in the p PIM set to 2 through 1 1 to reflect constant survivorship with time-dependent probability of capture. The 0(.) /?(t) model was then rerun using these PIMs to improve model fit. To evaluate apparent overdispersion of the data (c = 4.405), c was estimated using the median c approach and logistically regressed for c = 1 to 4.5 (4.5 represents a slightly higher value than the observed c of 4.405) at 5 intermediate points for each of 100 simulations (Cooch & White 2007). Results Radio-transmittered Jo ve^'. -Thirteen of 44 radio-transmittered White-winged Doves were known to have died during the monitoring period. All mortality was attributed to predation. Of these, eight were killed by domestic cats, two by raccoons, one by a hawk (Accipiter sp.), one by a domestic dog, and one in which the predator could not be determined. By the end of the monitoring period, 21 transmittered White-winged Doves had left the study area, the status of six doves was undetermined (a signal was received but the dove could not be visually located), and four were known to be alive. Estimated period survivorship for transmittered White-winged Doves over the 123 day period was 0.43 (95% Cl: 0.22-0.65) (Fig. 1). SMALL, BACCUS & SCHWERTNER 51 Figure 1. Kaplan-Meier survival curve for transmittered white-winged doves in Mason, Texas in 2006 with 95% confidence intervals. Following the protocol assuming constant survival across sample periods a daily survival can be determined as [(0.43)(123)]/365 = 0.145 and from this R = 6.12. The implication being that for the population to maintain a constant size, each adult would have to produce 6.12 young per year. Mark-recapture. -Sixty -nim untransmittered doves were recap¬ tured during the trapping period (49 once, 10 twice). Overall recapture rate was 14.5% and individual recapture rate was 12.4% (to date the highest recapture rate achieved for this species). The model for constant survival and time dependent probability of capture, with 12 days designated a trap occasion, was selected as the most appropriate model because it had the smallest AICc and its Akaike weight suggested it was 7 times more likely to fit the data than the other models. We used the median c approach to obtain an estimated c = 1.01 (SE = 0.018). Estimated pooled survivorship for untransmittered White¬ winged Doves was 0.16 for a 118 day period, an intuitively unrealistic value. However, none of the models fit the data sufficiently to provide results considered reliable. Goodness-of-fit tests consistently showed lack of model fit to the data from either 52 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 failure to reach numerical convergence or P- values < 0.0001. Consequently, mark-recapture calculations of survivorship were excluded from the analysis. Discussion Fundamental to the use of radio-transmitters is the assumption that transmitters do not adversely affect activity patterns and related parameters of individuals being monitored (White & Garrott 1990). The use of radio-transmitters in the study of avian species has been more problematic in meeting these assumptions than in other vertebrate taxa because of the added constraint of flight, which must be considered in such studies. Other than transmitter weight, method of attachment of radio-transmitters has been of particular interest (Gaunt et al. 1997), primarily as related to limiting movement and behavior (Hooge 1991). In recent years, studies of Mourning Doves and White-winged Doves have indicated that subcutaneously implanted transmitters with external antennas are a feasible, if not preferred, method for attachment of radio-transmitters (Schulz et al. 2001; Small et al. 2004b; Berdeen & Otis 2006). Previous studies evaluating radio-transmitter attachment methods for White-winged Doves in flight pens have been conducted. No differences were found in subcutaneously implanted doves versus controls for physiological parameters (Small et al. 2004b), hematological parameters (Small et al. 2005b) or time budget behavior analysis (Rosales 2000). In field studies with White-winged Doves, subcutaneously implanted radio-transmitters were used with a high degree of success (period mortality rates < 5%) over 120 days (Small et al. 2005a) and 96 days (Small et al. 2006b) in two other urban habitats. In Mason, Texas, recaptured White-winged Doves with subcutaneously implanted radio¬ transmitters showed no obvious signs of adverse reaction to the attachment technique (i.e., surgical procedure) (Small et al. 2007). SMALL, BACCUS & SCHWERTNER 53 Berdeen & Otis (2006) verified that subcutaneously implanted radio-transmitters had no substantial effect on Mourning Dove survival and met the assumption that radio-tagging did not affect survival. However, their study focused on hunter harvest as the primary source of mortality. Additionally, they had a higher survivorship for implanted individuals held 24 h prior to release compared to those released immediately. It is not realistic to assume constant annual survival from period survival rates. Because White- winged Doves are a game bird in Texas, mortality increases substantially during the 2-month dove hunting season each autumn. Therefore, annual survival estimates for this species, derived from a 123 d period during the spring and summer, most likely represent an overestimate of the true survival. However, this does not necessarily render the analysis meaningless. White-winged Doves typically lay two eggs per clutch. Based on estimates derived from this study, each pair would have to produce >6 successful 2-egg clutches per year to offset the high mortality rates of radio-transmittered birds in our study. The greatest documented number of clutch attempts by an individual White¬ winged Dove is four, two of which were successful (Schaefer et al. 2004). Based on previous studies, nest success, defined as at least one successful fledgling, is about 50%. Therefore, the mortality rates associated with transmittered birds appear to be unsustainable, suggesting that these rates are substantially higher than for normal (i.e., untransmittered) birds. Although previous studies using subcutaneously implanted radio-transmitters showed low mortality rates (<5%) (Small et al. 2005: 2006b), this study definitively had a minimum predation rate of 29.5% over the observational period. This suggests that in some circumstances implanted radio-transmitters may be detrimental to individual survivorship. Consequently, it is suggested that care be taken when designing radio-telemetry studies with this or similar species. 54 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Acknowledgments We wish to thank C. Lewis and J. Stay or for their assistanee in the field. Two anonymous reviewers provided eonstructive eomments on earlier drafts of the manuscript. This study was funded by the Texas Parks and Wildlife Department White- winged Dove Stamp Fund. Literature Cited Berdeen, J. B. & D. L. Otis. 2006. Effects of subcutaneous transmitter implants on mourning doves. Wildl. Soc. Bull., 34: 93-103. Burnham, K. P. & D. R. Anderson. 2002. Model selection and multimodel inference: a practical information-theoretic approach. Springer- Verlag, New York, New York, 496 pp. City-data.com [online]. 2005. Mason, Texas detailed profile. http://www.city-data.com/city/Mason-Texas.html (16 November 2006). Cooch, E. & G. White [online]. 2007. Program MARK: a gentle introduction. http://www.phidot.org/software/mark/docs/book/ (14 April 2007) Cottam, C. & J. B. Trefethen. 1968. Whitewings: the life history, status, and management of the white-winged dove. D. Van Nostrund, Princeton, NJ, 320 pp. Gaunt, A. S., L. W. Oring, K. P. Able, D. W. Anderson, L. F. Baptista, J. C. Barlow, & J. C. Wingfield. 1997. Guidelines to the use of wild birds in research. Ornithological Council, Washington, D.C., USA, 56 pp. George, R. R., E. Tomlinson, R. W. Engel-Wilson, G. L. Waggerman, & A. G. Spratt. 1994. White-winged dove. Pp. 29-50 in T. C. Tacha and C. E. Braun, editors. Migratory, shore and upland game bird management in North America. Allen Press, Lawrence, Kansas, 223 pp. George, R. R., G. L. Waggerman, D. M. McCarty, R. E. Tomlinson, D. Blankinship, & J. H. Dunks. 2000. Migration, harvest and population dynamics of White¬ winged Doves banded in Texas and northeastern Mexico, 1950-1978. Texas Parks Wildl. Dep., Austin, 49 pp. Gould, F. W., G. O. Hoffman, & C. A. Rechenthin. 1960. Vegetational areas of Texas. Texas A&M University and Texas Agricultural Experiment Station, College Station, Texas, USA, 492 pp. Guthery, F. S. 2002. The technology of bobwhite management. Iowa State University, Ames, lA, 224 pp. Guthery, F. S. & J. L. Lusk. 2004. Radiotelemetry studies: are we radio¬ handicapping northern bobwhites. Wildl. Soc. Bull., 32:194-201. Hayslette, S. E. & B. A. Hayslette. 1999. Late and Early Season Reproduction of Urban White-winged Doves in Southern Texas, Texas J. of Sci. 51(2): 173-180. Hooge, P. N. 1991. The effects of radio weight and harness on time budgets and movements of acorn woodpeckers. J. Field Omith., 62:230-238. SMALL, BACCUS & SCHWERTNER 55 Kaplan, E. L. & P. Meier. 1958. Nonparametric estimation from incomplete observations. J. Am. Statistical Assoc., 53:457^81. Pollock, K. H., S. R. Winterstein, C. M. Bunck, & P. D. Curtis. 1989. Survival analysis in telemetry studies: the staggered entry design. J. Wildl. Manage., 53:7- 15. Purdy, P. C. & R. E. Tomlinson. 1991. The eastern white-winged dove: factors influencing use and continuity of the resource. Pp. 225-265 in J. G. Robinson & K. H. Redford, editors. Neotropical wildlife use and conservation. University of Chicago Press, Chicago, Illinois, 538 pp. Reeves, H. M., A. D. Geis, & F. C. Kniffin. 1968. Mourning dove capture and banding. Special Scientific Report, No 117, U.S. Fish and Wildlife Service, Washington, D.C., 63 pp. Rosales, R. 2000. Effects of transmitter attachment techniques on captive White¬ winged Doves (Zenaida asiatica). MS Thesis. Sul Ross State University, Alpine, Texas, 68 pp. Schaefer, C. L. M. F. Small, J. T. Baccus, & R. D. Welch. 2004. First definitive record of more than two nesting attempts by wild White-winged Doves in a single breeding season. Texas J. of Sci., 56(2): 179-1 82. Schaefer, C. L., J. T. Baccus, M. F. Small, & R. Welch. 2005. Trapping and recapture rates for urban White-winged Doves in Waco, Texas. Bull, of Tex. Omith. Soc.,38(l):12-15. Schulz, J. H., A. J. Bermudez, J. D. Firman & Z. He. 2001. Comparison of radiotransmitter attachment techniques using captive mourning dove. Wildl. Soc. Bull. 29:771-782. Schulz, J. H., A. J. Bermudez, J. L. Tomlinson, J. D. Firman & Z. He. 1998. Effects of implanted radiotransmitters on captive mourning doves. J. of Wildl. Manage., 62:1451-1460. Schwertner, T. W. & K. Johnson. 2006. Using land cover to predict White-winged Dove occurrence and relative density in the Edwards Plateau. Pp. 98-102 in J. W. Cain, III & P. R. Krausman, editors. Managing wildlife in the southwest: new challenges for the 2U^ Century. Southwest Section of The Wildlife Society, Alpine, Texas, USA, 142 pp. Schwertner, T. W., H. A. Mathewson, J. A. Roberson, M. Small, & G. L. Waggerman. 2002. White-winged dove {Zenaida asiatica). Account No. 710 i« A. Poole & F. Gill, editors. The Birds of North America, Academy of Natural Sciences, Philadelphia, Pennsylvania, USA, and American Ornithologists’ Union, Washington D.C., 28 pp. Small, M. F., J. T. Baccus, & T. W. Schwertner. 2006a. Historic and current distribution and abundance of white-winged doves {Zenaida asiatica) in the United States. Tex. Omith. Soc., Occ. Pub., 6:1-23. Small, M. F., J. T. Baccus, & T. W. Schwertner. 2007. Observations of recaptured white-winged doves with subcutaneous radio-transmitter implants. Bull, of Tex. Omith. Soc., 40:29-31. Small, M. F., J. T. Baccus, & G. L. Waggerman. 2004a. Mobile anesthesia unit for implanting radio transmitters in birds in the field. Southwestern Nat., 49:279- 282. 56 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Small, M. F., J. T. Baccus, & G. L. Waggerman. 2006b. Breeding ecology of radio- tagged White-winged Doves {Zenaida asiatica) in the Coastal Bend region of Texas. Texas J. of Sci., 58(l):79-86. Small, M. F., C. L. Schaefer, J. T. Baccus, & J. A. Roberson. 2005a. Breeding ecology of a recently colonized urban white-winged dove population. Wilson Bull., 117:172-176. Small, M. F., J. T. Baccus, J. F. Mink, and J. A. Roberson. 2005b. Hematological responses in captive White-winged Doves {Zenaida asiatica) induced by various radiotransmitter attachments. Journal of Wildlife Diseases, 41(2):387-394. Small, M. F., R. Rosales, J. T. Baccus, F. W. Weckerly, D. N. Phalen, & J. A. Roberson. 2004b. A comparison of effects of radiotransmitter attachment techniques on captive white-winged doves. Wildl. Soc. Bull., 32:627-637. United States Geological Survey [online]. 1999. National land cover data: mapping procedures, (21 November 2006). White, G. C. & K. P. Burnham. 1999. Program MARK: Survival estimation from populations of marked animals. Bird Study 46 (Supplement): 120- 138. White, G. C. & R. A. Garrott. 1990. Analysis of wildlife radio-tracking data. Academic Press, San Diego, CA, 383 pp. MFS: msl253@txstate.edu TEXAS J. SCI. 60(1), FEBRUARY, 2008 57 GENERAL NOTES SOME OBSERVATIONS ON THE MATING BEHAVIOR OF THE GIANT WALKING STICK, MEGAPHASMA DENTRICUS (ORTHOPTERA: PHASMIDAE) Tara L Maginnis, Christina L Cool and Jana L Muniz Department of Biology, St. Edward’s University Austin, Texas 78704 Sexual dimorphism is a prominent feature throughout the animal kingdom, from birds to ungulates to arthropods (Blanekenhom et al. 2007). In many stiek inseets, sexual dimorphism is two-fold. First, the males may bare enlarged femoral spines, perhaps used as weapons in male-male eompetitions (Sivinski 1978; Broek 1999). Second, the males can be smaller than females, which may be a benefit to both males and females, as during mating, the male hangs on to the female, and a smaller male may allow the female to be more mobile for feeding and/or have more energy to invest in reproduction (Sivinski 1978). However, the idea that relatively small males have a higher fitness than large males is contradictory to a body of sexual selection theory that suggests that larger males, specifically in species that have weapons and engage in male-male competitions, often obtain more matings (Andersson 1994). The preliminary observations on the male-male and male-female interactions in the giant walking stick Megaphasma denticrus (Stal; also referred to as Megaphasma denticrum), while too few for statistical analyses, suggest that future investigations of the following hypotheses could be profitably pursued: (1) males compete with each other over access to females or resources, (2) large males are more likely to mate than small males, (3) males missing legs (autotomy) experience any mating disadvantages, (4) males prefer large females, and (5) females prefer small males. The giant walking stick is one of the largest phasmid species in North America and is found in the south central United States (Figure 1; Hebard 1943; Wilkings & Breland 1951). Both sexes 58 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Figure 1. A female (top) and male Megaphasma dentricus. have femoral spines on their mid and hind legs, but males have a single large spine and females a row of small spines. Both sexes are green and brown in eolor, and females are generally larger than males (females in this study ranged from « 105- 135 mm and males ranged from «90-125 mm). Females drop their eggs to the ground, and ean lay up to three eggs per hour and thirteen per day for several months. Like some other phasmids, males have a elasping organ - modified eerci used to wrap around a female’s abdomen during eopulation and/or mate guarding (Figure 2; Wilkins & Breland 1951; Sivinski 1978). In May 2007, 29 females and 30 males were eolleeted in Blanco County, Texas (mostly found on mesquite, Prosopis glandulosa Torr.). Individuals were brought to the lab, measured for body size (anterior of head to tip of abdomen), inspected for missing legs, and TEXAS J. SCI. 60(1), FEBRUARY, 2008 59 Figure 2. Clasping organ of a male (left) around a female. given a unique color band for identification (Figure 1). All animals were fed hackberry leaves ad libitum {Celtis laevigata Willdenow) and kept in cages in a large greenhouse. Groups of individuals (see Table 1) were placed in a 37.85 L fish tank and recorded for 12 h with a red light/ lamp and Sony Handycam DCR-SR42 («7:00 p.m. to «7:00 a.m.; like many insects, phasmids are thought to be unable to see red light). No individuals were used in observations more than once, and once used, individuals were put together in a large holding cage where their mating behaviors were recorded approximately every twelve hours. During filming, six categories of behaviors were noted: mating effort, mating, grappling, feeding, walking, and motionless. ‘Mating effort’ was defined as either a male simply hanging on a female (Figure 1) or a male attempting to wrap his clasping organ around the female. ‘Mating’ was defined as a male having his clasping organ wrapped around the female (Figure 2; actual copulation was not determined in this study because it was difficult to tell whether the genitalia were in physical contact or whether the male simply has his claspers around the female). ‘Grappling’ was defined as two individuals who engaged 60 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Table 1. Mating behavior in Megaphasma dentricus. Small males were shorter than 101.3 mm, and large males were longer than 115.7 mm. Similarly, small females were shorter than 110.2 mm, and large females were longer than 127.7 mm. All small and large pairs were at least 15 mm difference in size. Average males were «105 mm and average females were «1 17 mm. Individuals Trial Successful « Minutes until Time spent on mating effort by Mating Pair claspers attached Toser’ male (out of ^720min) Small cT 1 $ + Small cT 1 70.1% (505 min) Large cT Average ? 2 ? + Large cT <1 97.2% (700 min) 3 ? + Smaller 4 3.5% (25 min) 4 $ + Large cT 2 0.0% (0 min) 5 ? + Large cT 2 29.2% (210 min) d' * 1 ? + cT (-) 1 leg <1 47.9% (345 min) (-) 1 leg ? 2 ? + 2 3.5% (25 min) 3 ? + cf 2 91.0% (655 min) d * 1 $ + cT (-) 2 legs <2 39.6% (285 min) (-) 2 legs $ 2 $ + cT (-) 2 legs 1 41.7% (300 min) Small ? 1 cT + Large ? 2 N/A Large ? Average cT 2 cT + Small ? <1 N/A 3 cT + Small ? 3 N/A * Body size in males was controlled for in each of these experiments/trials (e.g., the male with all six legs and the male missing legs were within two mm of each other). in extensive touching of their legs and is considered to be mildly aggressive (Sivinski 1978). Males do not appear to compete with each other over food or females, as grappling behavior occurred less than 2% of the time (x = 10.9/720 min, SD ± 6.3). This was true for all trials involving females (listed in Table 1) and trials with only two males (e.g., small vs. small, large vs. large, small vs. large, and average vs. average - one trial each, not listed in Table 1, same filming methods as above). With or without a female present, males often came in contact without grappling, and simply walked over or around each other with no signs TEXAS J. SCI. 60(1), FEBRUARY, 2008 61 of aggression. In addition, they spent many minutes feeding in close proximity, and on several occasions, one male hung off the other. These preliminary observations suggest that mating is opportun¬ istic and on a ‘first-fmd, first-mate’ basis (Table 1). Large males were no more likely to find and successfully attach to females first, and males missing appendages appeared to be at no mating disadvantage. Females may show little preference for small males, and males no preference for large females; a male clasped to the first female he encountered, and remained attached for at least the next twelve hours, even if the other individual approached the mating pair. In each of the observations, a male had his claspers around a female in four minutes or less (most within two minutes), and he remained there for at least the next twelve hours. That is, there was no instance where a male successfully clasped around a female and then abandoned that female, even with no other male present. On many occasions within each of the 12h observations, the Toser’ male also got his claspers around the mating female, in which scenarios he attached anterior to the other male, further away from the female’s genitalia. This did not appear to be very stable, as this second male would repeatedly slip off the female. However, ‘second’ males spent up to 90% of their time attempting to mate with a paired male and female (x = 305/720 min, SD ± 253.7), often hanging or following the mating pair around the cage and/or swinging their abdomen around in an attempt to attach. On average, both males and females spent ^15% of their time feeding (x = 105.2/720 min, SD ± 71.7), and «85% of their time walking or motionless (x = 614.8/720 min, SD ± 71.7). The exceptions to this were single females; in the three observations with a male, a large female, and a small female, the unmated female spent up to 95% of her time feeding (x = 490/720 min, SD ± 190.8). Other interesting observations arose after the experimental animals were placed together in one large cage. First, two mating pairs were together for over 60 hours (but <72), presumably extensive mate guarding (Sivinski 1983; Brock 1999). Second, five males changed partners at least three times in a 36-hour period, and most males changed partners once every 24-36 hours. Moreover, their mate switching did not follow any pattern of preferring smaller or larger 62 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 individuals. Third, available males and females did not always form a mating pair (e.g., just because there was an available mate did not mean a mating pair formed). These observations provide more support that both males and females do not show strong preferences for body size or overall morphology of their mate. In conclusion, mating in Megaphasma dentricus appears to be opportunistic. Although there are strong adaptive hypotheses for both sexes related to their sexual dimorphism, our limited observations illuminated no apparent patterns of behavior involved in male-male competition or female choice. Other North American phasmids, such as Diapheromera veliei and D. covilleae, are different; larger males win contests over access to females and larger males mate longer than small females (Sivinski 1978). Perhaps the femoral spines function for crypsis and/or predator defense (Bedford 1978), and/or their sexual dimorphism is an evolutionary relic of other closely related species. Future work will reveal more details about the mating behavior seen in M dentricus, and if it is typical of other phasmids. Literature Cited Andersson, M. 1994. Sexual selection. Princeton University Press, Princeton, NJ, 624 pp. Bedford, G.O. 1978. Biology and ecology of the phasmatodea. Ann. Rev. Entomol., 23:125-49. Blanckenhom, W.U., A.F.G. Dixon, D. J. Fairbaim, M. W. Foellmer, P. Gibert, K. van der Linde, R. Meier, S. Nylin, S. Pitnick, C. Schoff, M. Signorelli, T. Teder & C. Wiklund. 2007. Proximate Causes of Rensch's Rule: Does Sexual Size Dimorphism in Arthropods Result from Sex Differences in Development Time? Am. Nat., 169(2):245-257. Brock, P. 1999. The amazing world of stick and leaf insects. The amateur entomologist, 26. Cravitz Printing Company Ltd., Brentwook, UK, 165 pp. Hebard, M. 1943. The dermaptera and orthoperous families blattidae, mantidae and phasmidae of Texas. Trans. Amer. Ent. Soc., 68:239-313. Sivinski, J. 1978. Intrasexual selection in the stick insects diapheromera veliei and D. covilleae and sexual dimorphism in the phasmatodea. Psyche, 85:395-405. Sivinski, J. 1983. Predation and sperm competition in the evolution of coupling durations, particularly in the stick insect Diapheromera veliei. Pp, 147-162, in Gwynne, D. T. and G. K. Morris (eds.), Orthoptera mating systems: Sexual competition in a diverse group of insects. Wesview Press, CO, 376 pp. Wilkins, O. P. & O. P. Breland. 1951. Notes on the giant walking stick Megaphasma denticrus (StM) (Orthoptera: Phasmida). Texas J. Science, 2:305-310. TLM at: taram@stedwards.edu TEXAS J. SCI. 60(1), FEBRUARY, 2008 63 FIRST REPORT OF OCHETOSOMA ANIARUM (DIGENEA: OCHETOSOMATIDAE) FROM THE BRAZOS WATER SNAKE, NERODIA HARTERI (SERPENTES: COLUBRIDAE), EnJ TEXAS, WITH A SUMMARY OF DEFINITIVE HOSTS OF THIS PARASITE Chris T. McAllister and Charles R. Bursey Department of Physieal & Life Sciences Chadron State College C hadron, Nebraska 69337 and Department of Biology Pennsylvania State University-Shenango Campus Sharon, Pennsylvania 16146 The Brazos water snake, Nerodia harteri, is a medium-sized colubrid endemic to the Brazos River system of 11 counties of northcentral Texas (Dorcas & Mendelson 1991; Conant & Collins 1998; Dixon 2000; Werler & Dixon 2000; Ernst & Ernst 2003). Discovered in 1936 in Palo Pinto County in the Brazos River, the species was later described by Trapido (1941). This snake fills a unique niche within this riverine habitat typified by fast-flowing rocky stream riffles of the upper Brazos River and two of its tributaries. Although not currently listed as federally threatened or endangered by the U.S. Fish and Wildlife Service, the species is considered threatened by the Texas Parks and Wildlife Department (TPWD 2007). In addition, N. harteri is globally listed G2 (NatureServe 2007) due to its restricted range and habitat requirements. Detailed information is available on various aspects of the natural history of this snake (Mecham 1983; Scott et al. 1989); however, very little is known about its parasites. McAllister & Upton (1989) reported three eimerians and a Cryptosporidium sp. from Brazos water snakes from Somervell and Palo Pinto counties, and Upton et al. (1989) further described a Cryptosporidium sp. in a single N. harteri from the latter county. The only previous report on helminth parasites was by Rossi & Rossi (2000) who reported Strongyloides sp. and unidentified filiarial worms from captive N. harteri. 64 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 On 18 July 1987 and again between 6 May 1988 and 29 July 1988, 10 juvenile and adult N. harteri (four males, six females; SVL range = 340-710 mm, mean ± \SD = 547.5 ± 113.4 mm) were collected by hand or tong from two sites on the Brazos River, one below Possum Kingdom Dam, 1 1.3 km SW Graford off FM 4, Palo Pinto County {n = 3, 32° 52.02’N, 98° 25.03’W), and the other 8.0 km S Glen Rose off FM 200, Somervell County {n = 7, 32° 16.16’N, 97° 39.5 1’W). Several of these snakes were the same specimens reported in McAllister & Upton (1989). Specimens were placed in individual collecting bags on ice and within 24 hr anesthetized with sodium pentobarbital (Nembutal®) overdose, and their gastrointestinal tract from their mouth to cloaca examined for helminth parasites. Trematodes were fixed in 10% neutral buffered formalin, transferred to 70% ethanol, stained with Semichon’s acetocarmine, and mounted in Canada Balsam. Voucher specimens of parasites are deposited in the United States National Parasite Collection (USNPC), Beltsville, Maryland, USA, as USNPC 84293. Snake voucher specimens are deposited in the Arkansas State University Museum, Herpetological Collection, State University, Arkansas, as ASUMZ 8420, 11221, 11762-11766, and the Cedar Valley College Collection (CVC), Lancaster, Texas, as 880520-18, 880708-6, and 880729-1. One of 10 (10%) N. harteri (655 mm SVL female, CVC 880708-6) collected from the Somervell County site on 8 July 1988 was found to be infected in its mouth with four specimens of the digenean trematode Ochetosoma aniarum. This was the only helminth found in this small sample of snakes. The parasite was originally described by Leidy (1890) as Distomum aniarum from the northern water snake, Nerodia sipedon, from Pennsylvania. In addition, Dubois & Mahon (1959) listed Renifer acetabularis, R. orula, R. natricis, R. texanus, and R. wardi as synonyms of O. aniarum. It would appear that Dubois & Mahon (1959) were unaware that Skrjabin & Antipin (1957) had previously transferred the above species of Renifer to Ochetosoma. The current arthors are in agreement with the latter synonymy although some disagreement currently exists (see Ernst & Ernst 2006). TEXAS J. SCI. 60(1), FEBRUARY, 2008 65 Table 1. North American definitive hosts of Ochetosoma aniarum. Variously reported as synonyms Neorenifer aniarum, Neorenifer acetabularis, Renifer acetabularis, R. orula, R. natricis, R. texanus, and R. wardi. Host scientific names follow Crother et al. (2000) and Crother et al. (2003). Family/Host Locale Reference Colubridae Coluber constrictor constrictor unknown Wallander(1968) C. constrictor flaviventris Nebraska Brooks (1979) C. constrictor foxii unknown Wallander(1968) Farancia abacura abacura unknown MacCallum(1921) Heterodon platirhinos Texas Harwood (1932) unknown Wallander(1968) Heterodon simus unknown Wallander(1968) Lampropeltis getula floridana Florida Parker (1941) unknown Wallander(1968) L. getula holbrooki Louisiana Rabalais (1969) Texas Dronen & Guidry ( 1 977) Nerodia cyclopion Florida Parker (1941) Illinois Dyer & Ballard (1991) Louisiana Rabalais (1969); Brooks (1979); Fontenot & Font (1996) Mississippi Byrd (1935) N. erythrogaster erythrogaster North Carolina Collins (1969) Tennessee Parker (1941) N. erythrogaster flavigaster Alabama Detterline et al. (1984) Arkansas (Lonoke County) previously unpublished Illinois Dyer (1999) Louisiana Rabalais (1969); Brooks (1979) Mississippi Byrd (1935) N. erythrogaster transversa Mexico (Nuevo Leon) Jimnez & Caballero (1975) Texas Curfman & Davidson (1966); previously unpublished N. fasciata confluens Louisiana Rabalais (1969); Brooks (1979); Fontenot & Font (1996) Texas Harwood (1932) N. fasciata fasciata Louisiana Rabalais (1969) N. fasciata pictiventris Florida Parker (1941) N. harteri Texas This study (new host record) N. rhombifer rhombifer Illinois Dyer ( 1 999) Kansas Crow (1913) Louisiana Rabalais (1969); Fontenot & Font (1996) Mississippi Byrd (1935); (1937) Tennessee Parker (1941) N. sipedon pleuralis Alabama Detterline et al. (1984) North Carolina Collins (1969) Texas Harwood (1932) N. sipedon sipedon Michigan Talbot (1934) Pennsylvania Leidy (1890) N. taxispilota Georgia Camp (1980) North Carolina Collins (1969) Seminatrix pygaea Florida Parker (1941) Viperidae Agkistrodon piscivorus leucostoma Louisiana Rabalais (1969) A. piscivorus piscivorus North Carolina Collins (1969) 66 THE TEXAS JOURNAL OE SCIENCE-VOL. 60, NO. 1, 2008 This parasite has previously been reported in two viperids and several colubrid snakes from various North American localities, including Texas (Table 1). From this summation it is obvious that most of these hosts are semiaquatic or aquatic species (primarily Nerodia spp.) that occasionally feed on frogs. In the life cycle, eggs containing miracidia hatch after being eaten by pulmonate snails {Physa spp.) and develop into daughter sporocysts. After leaving snails, cercariae penetrate and encyst in tadpoles of the genera Lithobates, Hyla and Pseudacris , which, when fed to snakes of the genus Nerodia, adult flukes occurred in the mouth and esophagus of definitive hosts 35 days later (Byrd 1935; Walker 1939; Schell 1985). Although N. harteri most often feeds on small fishes, primarily minnows (Tennant 1984; Werler & Dixon 2000) related neonate Concho water snakes, N. paucimaculata, has been reported to eat cricket frogs, Acris crepitans, by Greene et al. (1994). This may help explain the low prevalence of O. aniarum in N. harteri. Acknowledgments We thank B. D. Earle (Cedar Valley College) for assistance with collecting, owners of I. F. Anderson Farms (Lonoke, Arkansas) for allowing the senior author to collect on their properties, and the Arkansas Game and Fish Commission and Texas Parks and Wildlife Department for Scientific Collecting Permits nos. 775, and SPR-044-1 and SPR-0390-027, respectively, issued to C.T.M. Literature Cited Brooks, D. R. 1979. New records for amphibian and reptile trematodes. Proc. Helminthol. Soc. Washington, 46:286-289. Byrd, E. E. 1935. Life history studies of Reniferinae (Trematoda, Digenea) parasitic in Reptilia of the New Orleans area. Trans. American Microsc. Soc., 54:196-225. Byrd, E. E. 1937. A new trematode parasite, Renifer wardi n. sp., from the watersnake, Matrix rhombifera, from Columbus, Mississippi. J. Parasitol., 22:229-231. Camp, C. D. 1980. The helminth parasites for the brown water snake, Nerodia taxispilota, from Kinchafoonee Creek, Georgia. Proc. Helminthol. Soc. Washington, 47:276-277. Collins, R. F. 1969. The helminths of Matrix spp. and Agkistrodon piscivorus piscivorus (Reptilia: Ophidia) in eastern North Carolina. J. Elisha Mitchell Soc., 85:141-144. TEXAS J. SCI. 60(1), FEBRUARY, 2008 67 Conant, R. & J. T. Collins. 1998. A field guide to reptiles and amphibians of eastern and central North America, Edition, expanded. Houghton-Mifflin, Boston, Massachusetts, 616 pp. Crother, B. I. (ed.). 2000. Scientific and standard English names of amphibians and reptiles of North America north of Mexico, with comments regarding confidence in our understanding. SSAR Herp. Circ., 29:1-82. Crother, B. I., J. Boundy, J. A. Campbell, K. De Quieroz, D. Frost, D. M. Green, R. Highton, J. B. Iverson, R. W. McDiarmid, P. A. Meylan, T. W. Reeder, M. E. Seidel, J. W. Sites, Jr., S. G. Tilley & D. B. Wake, 2003. Scientific and standard English names of amphibians and reptiles of North America north of Mexico: Update. Herpetol. Rev., 34:196-203. Crow, H. E. 1913. Some trematodes of Kansas snakes. Kansas Univ. Sci. Bull., 7:125- 134. Curfman, B. E. & F. F. Davidson. 1966. Parasitic helminths of Matrix erythrogaster transversa (Hallowell), the blotched water snake, in the Waco, Texas area. Texas J. Sci., 18(3):121. Detterline, J. L., J. S. Jacob & W. E. Wilhelm. 1984. A comparison of helminth endoparasites in the cottonmouth {Agkistrodon piscivorus) and three species of water snakes {Nerodia). Trans. American Microsc. Soc,, 103:137-143. Dixon, J. R. 2000. Amphibians and reptiles of Texas: with keys, taxonomic synopses, bibliography, and distribution maps, 2"^* Edition. Texas A&M University Press, College Station, Texas, 421 pp. Dorcas, M. E. & J. R. Mendelson, III. 1991. Distributional notes on Nerodia harteri harteri in Parker and Palo Pinto counties, Texas. Herpetol. Rev. 22:1 17-1 18. Dronen, N. O. & E. V. Guidry. 1977. The use of selected ratios as an additional comparative tool in the systematics of some digenetic trematodes (Ochetosomatidae). Proc. Helminthol. Soc. Washington, 44:223-225. Dubois, G. & J. L. Mahon. 1959. Etude de quelques trematodes nord-americans (avec note sur la position systematique de Parorchis Nicoll, 1907 suivie d’une revision des generes Galactosomum Looss 1899 et Ochetosoma Braun 1901. Bull. Soc. Neuchatel Sc. Nat., 82:191-229. Dyer, W. G. 1999. Newly reported snake hosts for two ochetosomatid digeneans in Illinois. Trans. Illinois Acad. Sci., 92:243-245. Dyer, W. G. & S. R. Ballard. 1991. Ochetosoma aniarum (Leidy, 1891) Skrjabin and Antipin, 1957 (Trematoda: Plagiorchiidae) in Nerodia cyclopion (Dumeril, Bibron and Dumeril, 1854). Trans. Illinois Acad. Sci., 84:145-149. Ernst, C. H. & E. M. Ernst. 2003. Snakes of the United States and Canada. Smithsonian Books, Washington, D.C., 236 pp. Ernst, C. H. & E. M. Ernst. 2006. Synopsis of helminths endoparasitic in snakes of the United States and Canada. Soc. Stud. Amph. Rept. Herp. Circ., 34:1-86. Fontenot, L. W. & W. F. Font. 1996. Helminth parasites of four species of aquatic snakes from two habitats in southeastern Louisiana. J. Helminthol. Soc. Washington, 63:66-75. Greene, B. D., J. R. Dixon, J. M. Mueller, M. J. Whiting & O. W. Thornton. 1994. Feeding ecology of the Concho water snake, Nerodia harteri paucimaculata. J. Herpetol., 28:143-161. Harwood, P. D. 1932. The helminths parasitic in the Amphibia and Reptilia of Houston, Texas, and vicinity. Proc. U.S. Natl. Mus., 81:1-71. 68 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Jimnez, F. & E. Caballero. 1975. Reptilian trematodes in Nuevo Leon, Mexico. I. Ochetosoma chauhani sp. n. and O. aniarum (Leidy, 1871) Skrjabin et Antipin, 1957. Pp. 159-161, in Dr. B. S. Chauhan commemoration volume (K. K. Tiwari & C. B. Srivastava, eds.), Zool. Soc. India, Orissa, India. Leidy, J. 1890. Notices of Entozoa. Proc. Acad. Nat. Sci. Philadelphia, 42:410-418. MacCallum, G. A. 1921. Studies in helminthology: Part 1- Trematodes, Part 2- Cestodes, Part 3- Nematodes. Zoopathologica, 1:135-294. McAllister, C. T. & S. J. Upton. 1989. Coccidian parasites (Apicomplexa: Eimeriidae) of Nerodia spp. (Serpentes: Colubridae), with a description of a new species of Eimeria. J. Protozool., 36: 271-274. Mecham, J. S. 1983. Nerodia harteri. Cat. American Amph. Rept., 330.1-330.2. NatureServe 2007. 2007. NatureServe Explorer: An online encyclopedia of life [online] .Version 6.2. NatureServe, Arlington, Virginia. Available: http:www.natureserve.org/explorer [August 2007]. Parker, M. V. 1941. The trematode parasites from a collection of amphibians and reptiles. J. Tennessee Acad. Sci., 16:27-44. Rabalais, F. C. 1969. A checklist of trematodes from Louisiana snakes. Louisiana Acad. Sci., 32:18-20. Rossi, J. & R. Rossi. 2000. Comparison of growth, behavior, parasites and oral bacteria of Brazos water snakes, Nerodia harteri harteri, raised in an outdoor enclosure with related specimens raised indoors. Bull. Chicago Herpetol. Soc., 35: 221-228. Schell, S. C. 1985. Trematodes of North America north of Mexico. Univ. Press of Idaho, Moscow, Idaho, 263 pp. Scott, N. J., T. C. Maxwell, O. K. Thornton, L. A. Fitzgerald & J. W. Flury. 1989. Distribution, habitat, and future of Harter’s water snake, Nerodia harteri, in Texas. J. Herpetol., 23:373-389. Skrjabin, K. 1. & D. N. Antipin. 1957. Plagiorchioidea Dollfus, 1930. Pp. 453-598, in Trematodes of animals and man (K. 1. Skrjabin, ed.), Moskva 13:740 pp. Talbot, S. B. 1934. A description of four new trematodes of the subfamily Reniferinae with a discussion of the systematics of the subfamily. Trans. American Microsc. Soc., 53:40-56. Tennant, A. 1984. The Snakes of Texas. Texas Monthly Press, Austin, 561 pp. Trapido, H. 1941. A new species of Natrix from Texas. American Midi. Nat., 32: 473- 680. TPWD. 2007. Endangered and threatened amphibians and reptiles in Texas and the United States. Texas Parks and Wildlife Department, Austin, Texas, Available: http://www.tpwd.state.tx.us/huntwild/wild/species/endang/animals/reptiles_amphibia ns/ [June, 2007] Upton, S. J., C. T. McAllister, P. S. Freed & S. M. Barnard. 1989. Cryptosporidium spp. in wild and captive reptiles. J. Wildl. Dis., 25:20-30. Walker, J. H. 1939. Experimental studies on trematodes belonging to the subfamily Reniferinae. Trans. American Microsc. Soc., 58: 404-430. Wallander, R. W. 1968. The snake fluke Ochetosoma aniarum. Bull. Philadelphia Herp. Soc., 16:36-39. Werler, J. E. & J. R. Dixon. 2000. Texas snakes: identification, distribution, and natural history. University of Texas Press, Austin, Texas, 437 pp. CTM at: drctmcallister@aol.com TEXAS J. SCI. 60(1), FEBRUARY, 2008 69 INTRODUCTION OF THE BLUEFIN KILLIFISH {LUCANIA GOODEI) IN TEXAS Benny J. Callaway, Robert G. Fechhelm and Robert G. Howells* LGL Ecological Research Associates, Inc. 1410 Cavitt Street, Bryan, Texas 77801 and *BioStudies, 160 Bearskin Trail Kerrville, Texas 78028 In October 1997, E.E Dupont de Nemours and Company, Inc. constructed a 22.9“ha artificial wetland at a plant near Victoria, Victoria County, Texas, as part of a new biotreatment process of effluent water prior to its discharge into the Guadalupe River. As recently as 2001, the wetland had 10.6 ha of open water, bordered by extensive growths of emergent vegetation that had been planted on shallow shelves bordering several deep-water zones and on shallow flats separating the deep-water zones. Vegetation consisted of 24 species of marsh and water plants. Most were obtained from an onsite nursery, but three species (pickerelweed, Pontederia cordata; softstem bulrush, Scirpus validus; and giant cutgrass, Zizaniopois miliacea) were imported from a Elorida nursery (Duckworth-Cole, Inc. 2001). Eollowing completion of the planting of the aquatic vegetation, the Texas Parks and Wildlife Department (TPWD) seined a collection of fish from the Guadalupe River and released some of them into the wetland in late 1997 (Duckworth-Cole 1999). All were native species considered compatible with the wetland habitat, following acquisition of the property in 2004 by iNViSTA, S.a.r.L, the wetland became known as the INVISTA Wetland. Beginning in 1998, the wetland was sampled by seine and hoop net annually through 2002 and again in 2004 to determine the fate of the introduced fish stocks over time. When the wetland was first sampled in May 1998, collections included not only native fishes but also 20 small (26 to 40 mm in total length) killifish, tentatively 70 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 identified as the bluefin killifish (Lucania goodei) by Howells (2001). Prior to these collections, the introduced range of the bluefin killifish was believed restricted to Florida, southeastern Georgia, and southeastern Alabama, with introduced populations in North and South Carolina (Gilbert & Burgess 1980; Page & Burr 1991; Mettee et al. 1996; Fuller et al. 1999). This species is represented in the aquarium trade (e.g., Axelrod & Schultz 1971) and, as such, it is a candidate for introductions outside its range. In the wild, it inhabits vegetated ponds, lakes, and other still waters such as sloughs, pools and backwaters of streams. During spawning, adhesive eggs are deposited on vegetation or algae (Mettee et al. 1996). Giant cutgrass dominates the perimeter shores of the open-water cell where bluefin killifish were collected, and the species may have been introduced into the wetland as eggs attached to plants imported from Florida. By 2000, it became clear that the bluefin killifish was reproducing in the Invista Wetland. Fish increased in total length (TL) from 26-40 mm in spring 1998, to 30-52 mm in spring 1999. The appearance of numerous small fish 22-36 mm TL in spring 2000 indicated that successful spawning had occurred. Subsequent sampling periods (Table 1) demonstrates that the species is established and is successfully reproducing in the Invista Wetland. A representative male and female bluefin killifish were sent to the TPWD Heart of the Hills Fisheries Science Center (HOH), Ingram, Texas, for confirmation of the identification. Voucher specimens were placed in the Texas Cooperative Wildlife Collection (Catalogue number 10492.01), Texas A&M University, College Station, Texas. Additional specimens remain with LGL Ecological Research Associates, Inc., Bryan, Texas, and with the HOH. Water discharged from the Invista Wetland overflows a weir and drops vertically about 1 m into a large underground pipe. This pipe empties into an open channel that flows into holding ponds that also TEXAS J. SCI. 60(1), FEBRUARY, 2008 71 Table 1. Number of bluefin killifish (Lucania goodei) collected by 2-mm size intervals (total length) from the Invista Wetland, Victoria County, Texas. Length Interval (mm) 1998 1999 2000 2001 2004 22 1 3 24 2 2 26 3 6 3 28 1 15 8 30 2 1 23 1 6 32 4 14 1 5 34 5 1 12 1 10 36 3 8 13 2 7 38 1 23 10 1 15 40 1 36 11 2 18 42 58 7 1 16 44 68 11 1 2 46 75 11 1 3 48 56 8 1 1 50 23 6 1 52 6 2 1 54 1 56 1 Total 20 355 154 13 100 collect plant site runoff water and water discharged from an onsite cooling basin. The Invista Wetland is not accessible to fish from the holding ponds, but fish from the wetland can be transported into the holding ponds. In May 2002, 526 bluefin killifish were collected from the holding ponds. Water collected here is discharged through an underground pipe into a backwater slough of the Guadalupe River. Seining surveys in this slough conducted in May 2002 yielded 19 bluefin killifish. Bluefin killifish have not only established a reproducing population in the wetland, but, apparently, they have also dispersed from this site through the holding ponds into the Guadalupe River. 72 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 1, 2008 Literature Cited Axelrod, H. R. & L. P. Schultz. 1971. Handbook of tropical aquarium fishes. Tropical Fish. Hobbyist Publications, Neptune City, New Jersey, 718 pp. Duckworth-Cole, Inc. 1999. 1999 DuPont Wetland Assessment Report. Report for DuPont Wetland Assessment Team, E.I. DuPont De Nemours and Company, Inc. Victoria, Texas, 84 pp. + Append. Duckworth-Cole, Inc. 2001. 2001 DuPont Wetland Assessment Report. Report for DuPont Wetland Assessment Team, E.I. DuPont De Nemours and Company, Inc. Victoria, Texas, 142 pp. + Append. Fuller, P. L., L. G. Nico & J. W. Williams. 1999. Nonindigenous fishes introduced into the inland waters of the United States. Amer. Fish. Soc., Sp. Pub. 27, Bethesda. Maryland, 613 pp. Gilbert, C. R. & G. H. Burgess. 1980. Lucania goodei Jordan bluefin killifish. Pp. 534 in Atlas of North American Freshwater Fishes (D.S. Lee, CR. Gilbert, CH. Hocutt, RE. Jenkins, DE. McAllister, & JR. Stauffer, Jr., eds.). North Carolina State Museum of Natural History, Raleigh, North Carolina, 854 pp. Howells, R. G. 2001. Introduced non-native fishes and shellfishes in Texas waters: an updated list and discussion. Texas Parks and Wildlife Department, Data Series No. 188, Austin, Texas. 27 pp. Mettee, M.F., P.E. O’Neil & J.M. Pierson. 1996. Fishes of Alabama and the Mobile Basin. Oxmoor House, Birmingham, Alabama, 820 pp. Page, L.M., & B.M. Burr. 1991. A field guide to freshwater fishes. North America, North of Mexico. Houghton Mifflin Company, New York, 432 pp. Robinson, L. & J. Culbertson. 2005. A synoptic survey for non-indigenous ichthyofauna in selected tidal bayous of Galveston Bay. Texas Parks and Wildlife Department, Dickinson, Texas, 105 pp. Sterba, G. 1967. Freshwater fishes of the world. Pet Library, Ltd., New York, 877 pp. BJG at: bgallaway@lgl.com TEXAS J. OF SCI. 60(1), FEBRUARY, 2008 73 Manuscripts intended for publication in the Journal should follow these guidelines and be submitted in TRIPLICATE to: Dr. Frederick B. Stangl, Jr. TJS Manuscript Editor Department of Biology Midwestern State University Wichita Falls, Texas 76308 www.texasacademyofscience.org INSTRUCTIONS TO AUTHORS Scholarly manuscripts reporting original research results in any field of science or technology, including science education, will be considered for publication in The Texas Journal of Science. Prior to acceptance, each manuscript will be reviewed by both knowledgeable peers and the editorial staff. Authors are encouraged to suggest the names and addresses of two potential reviewers to the Manuscript Editor at the time of submission of their manuscript. No manuscript submitted to t\\Q Journal is to have been published or submitted elsewhere. Excess authorship is discouraged. 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No part of this publication may be reproduced, stored in a retrieval system or transmitted, in any form or by any means, electronic, mechanical, recording or otherwise, without the prior permission of the Managing Editor of the Texas Journal of Science. THE TEXAS JOURNAL OF SCIENCE Volume 60, No. 2 May, 2008 CONTENTS A Reexamination of Syngnathus affinis Gunter 1870, with Comparisons to Syngnathus scovelli (Evermann and Kendall 1 896) (Teleostei: Syngnathidae). By James Tolan . . . . . . . 83 Isolation and Characterization of Nitrogen-Fixing Bacteria Nodulating Phaseolus vulgaris in the Lower Rio Grande Valley of Texas. By Luis A. Materon and Thomas Eubanks..... . . . . . 97 Notable County Records for Rolling Plains Angiosperms of North Central Texas. By Allan D. Nelson and Jim R. Goetze . . . . . . . . . . . 113 Herpetofaunal Inventory of Camp Mabry, Austin, Texas: Community Composition in an Urban Landscape. By Adam W. Ferguson, Molly M. McDonough and Michael R. J. Forstner . 123 Novel AutoCAD Method for Performing Surface Energetics Analyses. By Les N. McDaniel, Nicholas A. Romero, George P. Cobb and Gopal Coimbatore . . . . . . . . . . 137 General Notes Reproductive Cycle of the Common Barking Gecko, Ptenopus garrulous (Squamata: Gekkonidae) from Southern Africa. By Stephen R. Goldberg . . . . . . . . . 151 Site Factors Affecting Pinyon- Juniper Occurrence in the Capulin Volcano Field of Northeastern New Mexico. By W. David Hacker and Jennifer L. Grosso . . . . . 157 THE TEXAS JOURNAL OF SCIENCE EDITORIAL STAFF Managing Editor: Ned E. Strenth, Angelo State University Manuscript Editor: Frederick B. Stangl, Jr., Midwestern State University Associate Editor for Botany: Janis K. Bush, The University of Texas at San Antonio Associate Editor for Chemistry: John R. Villarreal, The University of Texas-Pan American Associate Editor for Computer Science: Nelson Passos, Midwestern State University Associate Editor for Environmental Science: Thomas LaPoint, University of North Texas Associate Editor for Geology: Ernest L. Lundelius, University of Texas at Austin Associate Editor for Mathematics and Statistics: E. Donice McCune, Stephen F. Austin State University Associate Editor for Physics: Charles W. Myles, Texas Tech University Manuscripts intended for publication in the Journal should be submitted in TRIPLICATE to: Dr. Frederick B. Stangl, Jr. TJS Manuscript Editor Department of Biology Midwestern State University Wichita Falls, Texas 76308 frederick.stangl@mwsu.edu Scholarly papers reporting original research results in any field of science, technology or science education will be considered for publication in The Texas Journal of Science. Instructions to authors are published one or more times each year in the Journal on a space-available basis, and also are available on the Academy's homepage at: www.texasacademyofscience.org AFFILIATED ORGANIZATIONS American Association for the Advancement of Science, Texas Council of Elementary Science Texas Section, American Association of Physics Teachers Texas Section, Mathematical Association of America Texas Section, National Association of Geology Teachers Texas Society of Mammalogists TEXAS J. OF SCI. 60(2):83-96 MAY, 2008 A REEXAMINATION OF SYNGNATHUS AFFINIS GUNTER 1870, WITH COMPARISONS TO SYNGNATHUS SCOVELLI (EVERMANN AND KENDALL 1896) (TELEOSTEI: SYNGNATHIDAE) James Tolan Texas Parks and Wildlife Department, Coastal Fisheries 6300 Ocean Dr., NRC 2501, Corpus Christi, Texas 78412 Abstract.-The Texas pipefish (Syngnathus affinis) is currently known from only a small number of specimens collected from geographically isolated locations in the vicinity of Corpus Christi Bay, Texas. The general rarity of this species, coupled with the fact that no new collections have been recorded in over 30 years, has led numerous researchers to postulate that this taxon represents a marine species that has become extinct. To determine the status of S. affinis, a reexamination of all known collections was undertaken and these were compared to recent collections of S. scovelli, a common syngnathid ranging throughout the Gulf of Mexico. Multivariate statistical techniques were used to compare the morphometric and meristic characters currently used to differentiate syngnathids. Analysis of Similarity revealed that a low degree of separation exists between meristic characters of the two species, and an even lower degree of separation was found with the morphological values. Based on the results of this study, the taxonomic validity of the nominal designation S. affinis is questioned. Previous records of S. affinis most likely represent misidentified specimens of S. scovelli. Considerable confusion surrounds the status of the Texas pipefish, Syngnathus affinis Gunther 1870. This nominal species is based on a single specimen (Lot 233) purchased at Steven’s sales room (London) and registered simply as “Louisiana” and the questionable nature of the type-locality was indicated by Gunther’s failure to discuss the source of the holotype (Dawson 1982). Nearly all occurrences of this species have come from the Corpus Christi Bay area of Texas, with only one other subsequent collection from Louisiana (five specimens from Prien Lake; Fowler 1933). A total of 36 specimens have reported since 1926, with 20 of the 30 Corpus Christi Bay specimens taken with S. louisianae and S. scovelli in two seine collections at Fish Pass (Dawson 1982). A single specimen collected in 1970 (but subsequently lost) constitutes the collection from the Laguna de Terminos area (Dawson 1982). 84 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Early accounts of short-snouted pipefishes (snout length-to-head length ratios of 0.400 to 0.588) of the genus Syngnathus from the western Gulf of Mexico originally included only S. fuscus and S. scovelli (see Evermann & Kendall 1894:109 as Siphostoma fuscus\ Breder 1929; Jordan et al. 1930; Herald 1942; Hoese 1958). These two speeies were separated primarily by the total number of trunk rings (19-21, rarely 18 in S. fuscus\ 15-18, rarely 19 in S. scovelli) and dorsal fm rays (35-43 in S. fuscus; 27-35 in S. scovelli). Based on a reexamination of the holotype. Herald (1965) was the first to adopt the subspeeies designation of S. fuscus affinis (Reliet Northern Pipefish), although it was originally postulated that the holotype was a speeimen of S. fuscus from an erroneous locality. Later, Hoese & Moore (1977) reeognized S. fuscus affinis as a subspeeies of S. fuscus known only from Corpus Christ! Bay. Syngnathus fuscus fuscus is distributed along the Atlantic coast from Gulf of St. Lawrence in Canada southwards to northern Florida (Lazzari & Able 1990). Dawson (1982) dropped the subspecies designation, using the combination of trunk rings (modally 18), tail rings (33-34), and dorsal fm ray counts (37-41) to separate S. affinis from S. fuscus. Other characters presented by Dawson (1982) for distinguishing S. affinis from other western Atlantie eongeners inelude moderate snout length (averages 0.42 snout-to-head length ratio) and depth (averages 0.33 snout depth-to- length ratio) in addition to a narrower preorbital bone. By eontrast, Hubbs et al. (1994) incorporated all previous Gulf of Mexieo specimens of S. fuscus as S. affinis. This practice was followed by McEaehran & Feehhelm (1998), thereby eliminating the Gulf of Mexieo from the distributional range of S. fuscus. In order to resolve some of the eonfusion surrounding the identity and status of S. affinis, a reexamination of all known eolleetions of this speeies was conducted and these were eompared to reeent eolleetions of S. scovelli from the Corpus Christ! Bay area. In this paper, the species status of S. affinis is questioned based on a multivariate analysis of the meristie and morphological characters eurrently used to identify pipefishes. TOLAN 85 Materials and Methods Collections of Syngnathus affmis were obtained from museums (Academy of Natural Sciences [ANSP], British Museum of Natural History [BMNH], California Academy of Science [CAS], Field Museum of Natural History [FMNH], Gulf Coast Research Laboratory [GCRL], Texas A&I University [TAIC], Texas A&M University [TCWC], Texas Memorial Museum [TNHC], National Museum of Natural History [USNM], and Yale University [YPM]). Collections of S. scovelli were made by the author from every location in the Corpus Christi Bay area where S. affmis were previously recorded (Fig. 1). Each specimen was examined using the characteristics outlined in Dawson (1982), with the following measurements taken with dial calipers under a stereo-microscope (lOX): head length, snout length, snout depth, trunk depth, anal depth, pectoral fm length, and dorsal fm base length. Measurements were taken to the nearest 0.1 mm and expressed as a percentage of head length. Meristic counts included: trunk rings, tail rings, doral fm rays, dorsal fm rings (subdorsal rings) over the trunk, subdorsal rings over the tail, and total dorsal fm rings. Multivariate statistical techniques were used to compare the similarity of the meristic and morphological characters among the samples (i.e., individual pipefish specimens). Analyses were performed using Primer-E (Version 6.0) software (Clarke & Warwick 2001). Samples were standardized to account for scale differences in measurement units. A matrix of Euclidean distance similarities between samples was created. Significant differences in rank similarities between groups of samples were tested by Analysis of Similarity (ANOSIM). In the ANOSIM procedure, the probability of a priori groupings of samples is estimated by repeated permutations of the data (i.e., repeated random relabelling of samples in the matrix). Values of the R statistic can range from - 1 to 1 , although R will usually fall between 0 and 1 with R values > 0.4 indicating higher degrees of discrimination between groups. Similarities between the samples are graphically represented with non-metric multidimensional scaling (MDS) ordinations (Kruskal 86 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 97°20'0"W 96°40'0"W Fig. 1. Sampling locations in the vicinity of Corpus Christi Bay of Sygnathus affinis (o, open circles) ans S. sovelli (•, solid circles) collected for this study. 1964). Although outcomes of the ANOSIM are not dependant on MDS ordinations, the ordinations are presented here as they are a helpful way of visualizing patterns in the data. Stress values indicate how well the two-dimensional plot represents relationships among samples in the multidimensional space. Stress values <0.15 indicate a good fit. MDS ordinations may be arbitrarily rotated so axes are not labeled. Material examined. -Syngnathus affinis {N = 106). Numbers in parenthesis are number of specimens followed by size range; measurements are in mm SL. “Louisiana”, BMNH 1854.7.3.2 TOLAN 87 (holotype)(l, 144); Texas, Corpus Christi Bay and vicinity: CAS 39674 (1, 80); FMNH 40309 (2, 206, 218); GCRL 15252 (10, 55- 123); TAIC 3807 (as SJuscus 1, 123); TAIC 5307 (as SJuscus 72, 65-148); TNHC 27813 (9, 78-93); TNHC 28102 (as S. fuscus 1, 91); USNM 132675 (2, 117-174); Galveston Bay, TCWC 11633.01 (1, 109); Florida, Egmont Key, YPM 8724 (1, 89); Louisiana, Prien Lake, ANSP 55455 (5, 95-102). Syngnathus scovelli {N =170), uncataloged. Texas, Corpus Christi Bay and vicinity: Corpus Christi Bay (16, 74-118); Fish Pass (18, 72-120); Upper Laguna Madre (9, 75-105), Baffin Bay (17, 60-86); Aransas Bay (2, 61-85); Redfish Bay (3, 57-113). Results A total of 170 pipefish were collected by the author between June 2005 and November 2005 from the Corpus Christi Bay area. Field identifications resulted in the collection of 148 S. scovelli and 22 S. louisianae. Other collections provided by the Texas Parks and Wildlife Department (TPWD) Coastal Fisheries’ systematic bag seine sampling efforts in Upper Laguna Madre, Corpus Christi, and Aransas bays during this same time period resulted in an additional 19 Y scovelli and 3 S. louisianae specimens. No new specimens of S. affmis were collected during this study. Individuals of S. scovelli (n = 65) were randomly selected from this collection and used in the multivariate comparisons. ANOSIM results of the meristic characters (Table 1) revealed a relatively low degree of separation of the a priori designations {R = 0.363, P < 0.001). MDS configuration of the meristic characters is shown in Fig. 2. The most cohesive group of S. affmis samples fell on the left side of the MDS, with the majority of these samples constituting the Fish Pass (GCRL 15252) and the Upper Laguna Madre (TAIA 5307) collections. The remainder of the S. affmis collections from other Texas bays, as well as the Louisiana (ANSP 55455) and Florida (YPM 8724) specimens were scattered throughout the ordination among the S. scovelli collections. Based on the meristic information, the holotype was located in the middle Table 1, Counts and proportional measurements of the holotype and non-type specimens of Syngnathus qffinis (Junter 1870 with comparisons to S. scovelU. Morphometric characters expressed in percent of head length, except for SL in mm and snout depth-to» length ratio. 88 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 If s: 'C w U I S ^ o U a: 1 - O S 23 I I I I o <"4 -*0 C'l m cn o o o o o o o o o o o o o cn — « ^ O. "5^ ®0 Ch ‘ri lA o o o o o o Al QO c*^ fN. 0 0 or rr in fA €^1 T'l 0 d fd 0® *0 rr cd 0 d d d d d •— ro d CSC m r- «!$• »n m m m m cso *cr 0 0 m *n m IN 00 S S 0 d d d d d d d d ee a .£ g -s 3: « >> o *3 *9 S = *s « -H -C _ o o % s — ■ s 2 2 ’C g ;s C 3 3 O H H c/5 c/5 s TOLAN 89 Fig. 2. Non-metric multidimensional scaling ordination of the multivariate analysis (Euclidean distance measure) performed on the meristic features of Syngnathus affmis (°, open circles) and S. scovelli (•, solid circles) collections. S. affinis holotype identified by the star (*) symbol. Sample superscripts correspond to the following museum collections: 1-GCRL 15252; 2-CAS 39674; 3-USNM 132675; 4-ANSP 55455; 5-TNHC 27813; 6-TAIA 5307, TAIA 3807; 7-YPM 8724, 8-FMNH 40309; 9-TCWC-l 1633.01; 10-BMNH 1854.7.3.2. of the variability exhibited by the S. affinis collections. Propor¬ tional morphological measurements between the groups were less distinct, as shown in Fig. 3. While statistically significant {R = 0.123; P < 0.001), the morphologically-based R value between S. affinis and S. scovelli is negligibly small. The test reveals that the two ‘species’ probably do not have exactly the same proportional characteristics (the null hypothesis R = 0 can be rejected) and that these measurements are strongly overlapping and differ somewhat {R is closer to zero). Unlike the meristic characters, ordination of the morphological information failed to detect any cohesive groupings of the S. affinis samples. The holotype was again located within the range of variability encompassed by S. affinis. 90 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Fig 3. Non-metric multidimensional scaling ordination of the multivariate analysis (Euclidean distance measure) performed on the morphological features of Syngnathus affinis and S. scovelli collections. Sample superscripts correspond to the following museum collections: 1-GCRL 15252; 2-CAS 39674; 3-USNM 132675; 4-ANSP 55455; 5-TNHC 27813; 6-TAIA 5307, TAIA 3807; 7-YPM 8724, 8-FMNH 40309; 9-TCWC-l 1633.01; 10-BMNH 1854.7.3.2. Discussion Little or no literature is available describing the specific characteristics, physiology, or habitat requirements of S. affinis. Presumably, S. affinis shares similar habitat (shallow, nearshore submerged aquatic vegetation) and food requirements as S. scovelli (Joseph 1957; Huh & Kitting 1985; Holland & Boettcher 2005). Because few specimens have been reported since 1977, this species is currently described as among the rarest in the Gulf of Mexico and it is presumed to be extinct (Roberts & Hawkins 1999). Reasons for disappearance include habitat decline and degradation, caused by human-influenced fishing, shrimping, and dredging activities (Roberts & Hawkins 1999; Musick et al. 2000). While seagrass decline is cited as a specific reason for this demise, seagrass beds in the vicinity of Corpus Christi Fish Pass (the area of highest concentrations of S. affinis to date) have increased dramatically TOLAN 91 between 1974 and 1994 (Pulich & White 1997). Similar declines in populations of S. scovelli from comparable locations around the Corpus Christi Bay area have not been noted (bag seine collections 1982-2005; TPWD Coastal Fisheries database, Austin, Texas). While not currently listed as threatened or endangered by the National Marine Fisheries Service, the American Fisheries Society recommends listing the Distinct Population Segments of S. affinis as ‘endangered’ because of their “general overall rarity, endemic nature, and restricted geographic range” (Musick et al. 2000). Extensive sampling of the known localities around the Corpus Christi Bay area where S. affinis had previously been reported was unsuccessful. In the limited time frame used for this study, a total of 192 pipefishes were collected from nearshore habitats. Syngnathus scovelli made up the majority of these collections, revealing that this species is indeed the dominant syngnathid found in coastal regions of the Gulf of Mexico (Bolland & Boettcher 2005). Admittedly, the field sampling used in this study was limited in scope, concentrating primarily on the immediate vicinities of known locals of S. affinis populations. A spatially larger scope of sampling within the shallow, nearshore habitats is performed by TPWD, which conducts systematic bag seine sampling in each bay system along the Texas coast as a fishery-independent measure used to track the relative abundance and sizes of finfish (Martinez-Andrade et al. 2005). Since 1975, approximately 37,500 bag seine samples in shallow, nearshore habitats along the Texas coast have produced over 3,600 S. scovelli (TPWD Coastal Fisheries database, Austin, Texas). By stark contrast, no S. affinis have been recorded during these 30 years of sampling. This systematic monitoring of nearshore habitats has revealed that the highest densities of S. scovelli are found in Coastal Bend region of Texas (San Antonio, Aransas, Corpus Christi, and Upper Laguna Madre bay systems; see Fig. 1), and these areas of abundant populations of S. scovelli are generally the same areas that were sampled extensively for this study. Given 92 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 the general overlap in the meristic characters between S. affinis and S. scovelli as outlined in this paper, it is quite possible that field identifications of S. scovelli by TWPD staff could have erroneously included at least some S. affinis in the thousands of individuals reported. Previous taxonomic keys (Parker et al. 1972; Hoese & Moore 1977; Murdy 1983) recognized S. fuscus as the only other short-snouted pipefish in the genus Syngnathus from Texas waters, yet S. fuscus was similarly absent from TPWD surveys during the 1975-2005 period. Again, misidentifications could account for this apparent absence of S. fuscus from TPWD surveys of the Texas coast. With no recent collections of S. affinis available for investigation, investigations of its specific status rest on existing museum specimens. This study did locate additional museum specimens {n = 74) of S. affinis not listed in Dawson (1982), and these were included in the present analysis. These new records call into question the limited distributional range of S. affinis, with this ‘species’ now recorded from around the northern Gulf of Mexico (1 from near Galveston Bay in Texas; 72 from the Upper Laguna Madre in Texas, and 1 from Egmont Key in Florida). This new range is fully encompassed by the known range of S. scovelli (Dawson 1982). The ANOSIM tests used for this study revealed that for the meristic information, a relatively low degree of separation was found between S. affinis and S. scovelli (Table 1). Early works describing S. affinis were primarily based on meristic differences found in the short-snouted pipefishes from the western Gulf of Mexico (Herald 1965; Parker et al. 1972), yet those works recognized that only a few individuals had ever been collected. The present analysis also found differences in the mean numbers of dorsal fm rays, trunk rings and tail rings, as well as trunk and tail subdorsal rings, although the ranges of these counts always overlapped, in some cases greatly, between the two nominal taxa. While the meristic-based ANOSIM was significant, this low of an R TOLAN 93 value is indicative of a high degree of overlap in the a priori groupings. This can happen when the number of replicates is large for the groups, thus giving a very large number of possible permutations. Biologically trivial differences can still be statistically significant when power is large. More importantly, the ANOSIM procedure based on the morphometric values failed to detect any consistent pattern of differences between the two forms. While the stress value calculated for the morphometric data was larger than the meristic configuration (0.12 vs. 0.06, see Fig. 2), the ordination of the morphological information still provides a potentially useful 2-dimensional picture (below the 0.15 level, see Clarke & Warwick 2001). Scaled as a function of head length, average ratios of snout depth, pectoral fm length, and dorsal fm base length were higher in S. affinis, whereas snout length, trunk depth, and anal depth were equivalent with S. scovelli. Each of these characters used in this analysis varied between the two forms by no more than, at most, the standard deviation of the average ratios. This lack of divergence can clearly be seen in the MDS ordination (Fig. 3). Given the plasticity of meristic characters within western Atlantic species of Syngnathus (e.g., tail rings ranging from 28 - 34 in S. scovelli and 32-35 in S. affinis\ dorsal fm rays ranging from 25 - 37 in iS. scovelli and 35 - 41 in 5*. affinis\ see Tables XL, XLI, XLII, and XLIII in Dawson 1982), the variability and gradients of these traits suggests that all the material examined for this study represent different phenotypes of S. scovelli. Individuals currently identified as S. affinis, on the basis of higher counts of trunk rings, tail rings, and dorsal fins rays, most likely represent individuals at the upper limits of these features currently seen in western Gulf of Mexico populations of S. scovelli. Based on the multivariate techniques used for this study, there appears to be little justification for recognizing S. affinis and S. scovelli as distinct species as the former is shown herein to be indistinct from the latter. 94 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Before invalidating S. affinis as a nominal taxon, extensive field work must be eondueted in the western Gulf of Mexieo to doeument that there is indeed only a single speeimen of short¬ snouted Syngnathus within the area. These eolleetions should undoubtedly eneompass a mueh greater temporal seale than the six months used for this study. Given the uneertainty in the type loeality for S. affinis, “Louisiana” is eertainly within the geographie range of S. scovelli. Morphometrie and meristie eharaeters of the holotype of S. affinis all lay within the range of variation found in S. scovelli. If invalidation of S. affinis ultimately proves to be the proper aetion, the Prineiple of Priority (International Commission of Zoologieal Nomenelature (ICZN) 2000) states that the valid name of any taxa be the oldest available name applied to it (in this ease, S. affinis is senior to S. scovelli). While S. affinis is senior, the Prineiple of Priority further states that it is not intended to be used to upset a long-aeeepted name in its aeeustomed meaning by the introduetion of a name that is its senior synonym or homonym (ICZN 2000: Artiele 23.2). Therefore, if further researeh eorroborates the present hypothesis of invalidating .S', affinis, all short-snouted pipefishes from the western Gulf of Mexieo should be referred to as S. scovelli. Acknowledgements I would like to thank the staffs of eaeh museum for allowing me permission to examine speeimens in their eare and for their prompt response to inquires and loan requests for the few existing eolleetions of the Texas pipefish. Patriek Campbell (BMNH) provided the morphometrie data from the holotype. This projeet was never explieitly funded by researeh grants, but I gratefully aeknowledge the eontinued support of Sportfish Restoration Funds, without whieh the time for data synthesis and interpretation would not have been possible. I would like to dedicate this paper to the memory of B. A. Thompson, Coastal Fisheries Institute, Louisiana State University, with whom my initial conversations about the origins of the Texas pipefish and the Prien Lake collections provided the original impetus for this work. TOLAN 95 Literature Cited Bolland, J. & A. Boettcher. 2005. Population structure and reproductive characteristics of the Gulf Pipefish, Syngnathus scovelli, in Mobile Bay, Alabama. Estuaries, 28(6):957-965. Breder, C. M., Jr. 1929. Field Book of Marine Fishes of the Atlantic Coast from Labrador to Texas. G.P. Putman’s Sons, New York: xxxvii + 322 p. Clarke, K. R. & R. M. Warwick. 2001. Change in marine communities: an approach to statistical analysis and interpretation, 2nd edition. PRIMER-E: Plymouth, 38 pp. Dawson, C. E. 1982. Fishes of the Western North Atlantic. Sears Foundation for Marine Research No. 1, Part 8. New Haven, Connecticut. 198 pp. Evermann, B. W. & W. C. Kendall. 1894. The fishes of Texas and the Rio Grande Basin, considering chiefly with reference to their geographic distribution. Bull. United States Fish Comm., 12:57-126. Fowler, H. W. 1933. Notes on Louisiana fishes. Proc. Biol. Soc. Washington, 46:57-64. Herald, E. S. 1942. Three new pipefishes from the Atlantic coasts of North and South America; with a key to the Atlantic American species. Stanford Ichthy. Bull.,2(4):125-134. Herald, E. S. 1965. Studies on the Atlantic American pipefishes with description of new species. Proc. California Acad. Sci. 4th Series, 32(12):363-375. Hoese, H. D. 1958. A partially annotated checklist of the marine fishes of Texas. Pub. Inst. Mar. Sci., 5:312-352. Hoese, H. D. & R. H. Moore. 1977. Fishes of the Gulf of Mexico; Texas, Louisiana, and adjacent waters. Texas A&M Univ. Press. College Station, Texas, 327 pp. Hubbs, C., J. D. McEachran, & C. R. Smith. 1994. Freshwater and Marine Fishes of Texas and the Northwestern Gulf of Mexico. The Texas System of Natural Laboratories, Inc. Series No. FTX/NWGM-94. Austin, Texas, 270 pp. Huh, S. H. & C. L. Kitting. 1985. Trophic relationships among concentrated populations of small fishes in seagrass meadows. J. Exp. Mar. Biol. EcoL, 92(l):29-43. International Commission of Zoological Nomenclature. 2000. International Code of Zoological Nomenclature, Fourth Edition. ISBN 0 85301 006 4. The Natural History Museum, London. Available online: http://www.iczn.org/iczn/index.jsp. Jordan, D. S., B. W. Evermann, & H. W. Clark. 1930. Check list of the fishes and fishlike vertebrates of North and Middle America north of the northern boundary of Venezuela and Columbia. Report of the United States Commissioner of Fisheries for 1928, Part II, 670 pp. Joseph, E. B. 1957. A study of the systematics and life history of the Gulf pipefish, Syngnathus scovelli (Evermann and Kendall). Ph.D. thesis. Florida State University, 105 pp. Kruskal, J. B. 1964. Multidimensional scaling by optimizing goodness of fit to a non-metric hypothesis. Psychometrika, 29:1-27. 96 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Lazzari, M. A. & K. W. Able. 1990. Northern pipefish, Syngnathus fuscus, occurrenees over the Mid-Atlantie Bight continental shelf: evidence of seasonal migration. Environ. Biol. Fish., 27:177-185. Martinez-Andrade, F., P. Campbell, & B. Fuls. 2005. Trends in relative abundance and size of selected fmfishes and shellfishes along the Texas Coast: November 1975-December 2003. Management Data Series No. 232. Texas Parks and Wildlife Department, Coastal Fisheries Division. Austin, Texas, 140 pp. McEachran, J. D. & J. D. Fechhelm. 1998. Fishes of the Gulf of Mexico. Vol. 1 - Myxniformes to Gasterosteiformes. Univ. of Texas Press, Austin, 1 1 12 pp. Murdy, E. O. 1983. Saltwater Fishes of Texas: A Dichotomous Key. Texas A&M University Sea Grant College Program, 220 pp. Musick, J. A., M. M. Harbin, S. A. Berkeley, G. H. Burgess, A. M. Eklund, L. Findley, R. G. Gilmore, J. T. Golden, D. S. Ha, G. R. Huntsman, J. C. McGovern, S. J. Parker, S. G. Poss, E. Sala, T. W. Schmidt, G. R. Sedberry, H. Weeks, & S. G. Wright. 2000. Marine, Estuarine, and Diadromous Fish Stocks at Risk of Extinction in North America (Exclusive of Pacific Salmonids). Fisheries, 25(ll):6-30. Parker, J. C. (ed.), B. J. Galloway, & D. Moore. 1972. Key to the Estuarine and Marine Fishes of Texas. Texas A&M University, 177 pp. Pulich, W., Jr. & W. A. White. 1997. Current status and historical trends of seagrass in the Corpus Christi Bay National Estuary Program area. Corpus Christ! Bay National Estuary Program Publication No. 20. Texas Natural Resource Conservation Commission, Austin, Texas, 131 pp. Roberts, C. M., J. P. Hawkins, N. Chapman, V. Clarke, A. V. Morris, R. Miller, & A. Richards. 1998. The threatened status of marine species. A Report to the World Conservation Union (lUCN), Species Survival Commission, and Center for Marine Conservation, Washington, D.C, 10 pp. Roberts, C. M. & J. P. Hawkins. 1999. Extinction risk in the sea. Trends Ecol. EvoL, 14(6):241-246. JT at: James. Tolan@tpwd.state.tx.us TEXAS J. OF SCI. 60(2):97-l 12 MAY, 2008 ISOLATION AND CHARACTERIZATION OF NITROGEN-FIXING BACTERIA NODULATING PHASEOLUS VULGARIS IN THE LOWER RIO GRANDE VAELEY OF TEXAS Luis A. Materon and Thomas Eubanks Department of Biology, University ofTexas-Pan American 1201 West University Drive, Edinburg, Texas 78541 Abstract. -To establish a highly effective symbiosis between a legume plant and its bacterial symbiont it is essential to develop a successful program for the selection of superior strains to maximize production through N2 fixation. Nitrogen-fixing bacteria from representative soil series of the Lower Rio Grande Valley region of southern Texas were examined for their symbiotic association with common ‘Pinto’ bean {Phaseolus vulgaris L.). PCR-RFLP analyses of genes associated with nitrogen fixation indicated that the bacterial isolates belong to the recently described Rhizobium etli and none were identified as R. tropici. Examination of symbiotic responses of the rhizobial isolates under controlled, aseptic conditions, indicated that 20% produced effective nodula-tion, 35% partially effective, and the remaining 45% produced parasitic or ineffective nodulation on the host legume. Strains UTPA-P 66, P82 and P80, each isolated from a different soil series, accounted for maximum plant response and nitrogen fixation activity. Although nodule morphology and size markedly varied among isolates, those with the highest tissue mass were associated with the highest production of shoot biomass. Electron micrographs of cross sections of 2-week old nodules indicated that effective bacterial cells were rod shaped, and more abundant than those enclosed in the peribacteroid membranes of ineffective nodules. To ensure adequate nodulation and N2-fixation, it is necessary to inoculate beans with locally isolated nitrogen-fixing bacteria, regardless of any previous history of bean cultivation. The inoculation of legume seeds with soil nitrogen-fixing bacteria (referred generically as ‘rhizobia’ or ‘rhizobial cells’) is a well-established technology of major economic and environmental importance. Natural populations of rhizobia can establish symbio¬ tic associations with legumes. The success of the symbiosis, which is reflected by enhanced yield, depends largely on the performance of the rhizobial strains (Materon & Zibilske 2001). When needed, the symbiotic association could be optimized through seed inoculation (Materon & Ryan 1996). A well-nodulated common bean {Phaseolus vulgaris) crop can fix up to 100 kg of N2 per ha annually, which is equivalent to 208.3 kg of urea fertilizer (Somasegaran & Hoben 1994). This legume is cultivated in areas 98 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 totaling 20,550 ha in the state of Texas (Smith & Anciso 2005). Most of this acreage is dedicated for commercial or processing purposes and approximately 60% of it is located in the Texas Winter Garden and in the High Plains region. The remaining production is obtained from small farms located in the southern part of the state (Smith & Anciso 2005). Unfortunately, according to many researchers (Graham 1981; Graham et al. 2003), inoculation with introduced inoculant rhizobial strains does not always lead to an abundant effective nodulation to secure high rates of nitrogen fixation and seed production (Graham 1981; Graham et al. 2002; Materon 1991; Materon & Zibilske 2001; Mrabet et al. 2005). This fact could be related to the promiscuity observed in P, vulgaris (cf Michiels 1998) or to other limiting factors, like the high rate of nitrogen fertilizer used in intensive agriculture, which is particularly detrimental for this legume species (Materon & Ryan 1996). In addition, this grain legume species also has a reputation for weakness in its ability to nodulate and efficiently fix nitrogen in symbiosis (Graham 1981; Graham & Halliday 1987; Graham et al. 2003). The diversity of rhizobial species that are able to nodulate P. vulgaris in relation to symbiotic effectiveness has been recently reported by various researchers (Amarger et al. 1997; Bal et al. 1982; Mostasso et al. 2002; Mrabet et al. 2005). A clearer knowledge of indigenous nitrogen-fixing bacteria is thus essential for the understanding of the ecological consequences of native biodiversity for legume production (Tamimi 2002). Information is virtually lacking on the symbiotic properties of indigenous populations of rhizobia and their capacity to nodulate and fix nitrogen when in symbiosis with locally-adapted cultivars of Phaseolus and other legume species grown in southern Texas. Other important characteristics of selected elite strains include their ability to compete and survive in the inoculant carrier and in soil, to colonize the rhizosphere, and to migrate in soil (Materon 1991; MATERON & EUBANKS 99 Materon & Zibilske 2001; Tamini 2002). It is expected that further research on rhizobial ecology will illustrate the relationship between population size in the rhizosphere and nodulation. Local farmers should avoid the application of nitrogen fertilizers due to the inefficient nitrogen fixation capacity of the indigenous and introduced inoculant strains. In the present study, a wide variety of agricultural soils were sampled in southern Texas to identify and evaluate the presence of any of five rhizobial species known to nodulate P. vulgaris. Materials and Methods Soil sampling procedures -Soil samples were collected from 83 agricultural sites representing the different soil types of the region. The sites were selected using soil survey maps of four counties (Hidalgo, Cameron, Willacy and Starr) located in the lower Rio Grande Valley of Texas. The Hidalgo soil series is classified as fine-loamy, mixed, active, hyperthermic typic Calciustoll; the Willacy as fine-loamy, mixed, super-active, hyperthermic Udic Argiustolls; the Cameron as Sodic Haplusterts; and, the Starr soil series as fme-loamy, mixed, semi-active, thermic Fluventic Dystrudepts (USDA, 2007). These soils are mildly to moderately alkaline with pH values ranging from 7 to 8, and very low soil organic matter contents ranging from 0.1 to 0.8 % organic carbon. The total area covered by these counties is 8,859 km^. At each sampling site, three soil samples were taken at 0, 5 and 10 m within a linear transect. Approximately 120 g of soil was collected from each point from a depth of 5-20 cm using a hand shovel. Samples were mixed and placed inside a plastic container and kept in a box away from sunlight. To prevent cross contamination, the shovel was sprayed in between sampling with a wash bottle containing a 50% Clorox™ (commercial hypochlorite bleach) solution, and rinsed with water three times (Beck et al. 1993). Elevation and air temperature were recorded as well as the longitude/latitude coordinates of the sampling site corresponding to the center of the transect using a manual Magellan Map 410 Global 100 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Positioning System (GPS) receiver. Samples were kept refrigerated at 3°C before analyses. The most probable number technique (MPN) procedure was applied for the calculation of number of nitrogen-fixing bacteria per gram using Phaseoulus vulgaris L. as a trap species (Vincent 1970). Inoculation and cultivation of bean seedlings. -Tmy^ containing layers of Sunshine™ soil mix potting medium made of composted bark, peanut hulls, Canadian sphagnum peat moss, perlite, and a wetting agent, were sterilized at 120°C, 0.1 MPa for 1 h. Plastic pots with a diameter of 10 cm were then filled to 70% capacity with the sterile potting medium. Seeds of P. vulgaris L. cv. ‘Pinto’ were surface sterilized by exposing them to 95% ethanol for 1 min followed by immersion in a 5.25% (vol/vol) sodium hypochlorite (NaOCl) solution for 1 min and then repeatedly rinsed in deionized sterile water (Beck et al. 1993; Vincent 1970). Two seeds were aseptically planted in each pot within a hole containing two spoonfuls of soil sample that acted as the inoculant with the intention to induce colonization and infection of the seedling root hairs. The inoculated seeds were then covered with more sterile potting soil. Pots were covered with saran wrap and distributed on tables Inside a greenhouse, located at the USD A Kika de la Garza Center for Subtropical Research in Weslaco (Texas), equipped for ambient temperature control and deionized water for irrigation purposes. The average temperature throughout the experiments was 21°C with a maximum of 23.6°C and a minimum of 15.6°C. Uninoculated controls included sixteen pots each with seeds and no soil inoculum. Aseptic handling of pots and instruments were meticulously practiced to avoid any cross contamination. Plants were watered once a day by an automatic sprinkler system using deionized water. Additional pots were set aside for early harvesting of effective and ineffective nodule samples for electron microscope observation. Plants were harvested after a 30-day period of growth MATERON & EUBANKS 101 for observation of patterns of nodulation and isolation of rhizobia from nodules. Isolation of rhizobia from trap legume host nodules -Koois were gently removed from pots and rinsed with running tap water to remove soil and debris. Three nodules were carefully detached from the crown area of the primary root. Nodules were immersed into a beaker with 95% ethanol for 10 sec, and then transferred into another beaker with a 2.5% (vol/vol) sodium hypochlorite (NaOCl) solution for 30 sec. Nodules were then rinsed five times in sterile deionized water to remove any excess of chlorine (Beck et al. 1993; Vincent 1970). Each nodule was crushed with a tip of a flamed broad tip forceps and its contents deposited onto the surface of a yeast-extract mannitol (YM) agar plate. Inoculated plates were incubated overnight at 37°C. Once bacterial growth was detected, a loopful of the confluent growth was streaked onto the surface of another YM agar plate to obtain single colonies. Samples of each isolate were examined microscopically using the Gram’s stain technique. Nitrogen-fixing bacteria specific to Phaseolus are rod shaped and Gram negative (Vincent 1970). All the presumptive isolates were grown aerobically on YM agar slants and stored at 4°C for further examination. Once tested for their symbiotic ability to produce N2-fixing nodules, isolates were assigned a collection name with the prefix UTPA-P followed by its respective catalog number as a suffix. To prevent genetic variation and loss of effectiveness traits, the bacterial cultures were lyophilized and kept in small vials at 4°C (Beck et al 1993; Somasegaran & Hoben 1994). Symbiotic evaluation of rhizobial cultures previously isolated from nodules -A total of 690 pots of 10.2 cm diameter were filled with coarse grade vermiculite that had been previously rinsed, drained with deionized water and sterilized using an autoclave set at 121°C and 0.1 mPa for 60 min. Two P. vulgaris L. cv. ‘Pinto’ bean seeds were surface sterilized (see procedure above) and planted in 102 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 each pot. The solutions of each of the bacterial cultures, grown at 27°C, had been adjusted by turbidity and plate count to provide a uniform density of approximately 10^ bacteria per ml (Beck et al. 1993; Somasegaran & Hoben 1994). One ml of the bacterial suspension was added to inoculate each of the seeds. Pots were then covered with a polyvinyl chloride plastic wrap to prevent any cross contamination during the early growth stage of the seedlings. After five days of growth an extra cover of plastic wrap was used to reseal around the stem of the plant. Pots were arranged in 23 groups of 30 replicated plants per isolate. Each group was split into three groups and randomly assigned a permanent position in the greenhouse. The inoculated treatments included an isolate from a commercial peat-based inoculant obtained from Becker Underwood (Ames, Iowa), labeled as strain 132, for comparison and contrast with the isolates under study. Uninoculated controls were not supplemented with nitrogen fertilizer, received no rhizobia, and were hydrated only with deionized water. Uninoculated nitrogen- fertilized controls were irrigated with N-free Hoagland solution supplemented with 5 mM KNO3 (Beck et al. 1993). All pots were regularly weighed and adjusted to a weight of 480 g by carefully adding N-free Hoagland solution. Plants were harvested after 30 days of growth. The rooting medium was carefully removed under a gentle stream of water so that not to detach nodules. For each plant, numbers and nodule distribution, as well as total nodule mass, were graded and recorded. Other plant parameters such as number of leaves and flowers were recorded. Means of dry shoot mass were used to measure plant response to inoculation for comparison with the nitrogen-deficient dry shoot mass mean of the uninoculated plants and referred to as the ‘symbiotic index’ (Beck et al. 1993; Vincent 1970). Genotypic characterization of rhizobial -Advanced molecular techniques allows a more accurate and precise determination of phylogenetic relationships among species. For this reason, PCR-RFLP analyses of symbiotic and genomic genes were used to determine the taxonomic position of twenty-one of the MATERON & EUBANKS 103 rhizobial strains able to nodulate P. vulgaris. Three restriction endonucleases, Hinfi, Rsal, and Alu\ were used in this study to digest the PCR amplification products for nodh, nijW, and GSII genes from all isolates representing the different locations sampled in the Lower Rio Grande Valley. Oligonucleotides used as primers included GSII--1&4 (Turner & Young 2000) for sequencing, nodA- 1&2 (Zhang et al. 2000) for the amplification of NodA, and nifH-l (Hau^a et al. 1998), and nifH-2 (Early et al. 1992) for the amplification of the nifH gene. RFLP products were run on 1.5% agarose gels, stained with ethidium bromide, and patterns were scored manually. Representatives of each of the PCR-RFLP types were sequenced using GSII PCR products. Purification using QIA quick PCR Purification Kit (Qiagen Hamburg GmbH) was performed following the manufacturer’s instructions. Sequencing was done for each product in both directions, and analyzed using Clustal-X (Thompson et al 1997) and BLAST analysis. The Basic Local Alignment Search Tool (BLAST) was used to identify regions of local similarity between sequences. Preparation of nodule tissues for transmission electron micros copy. -CX^diW 14-day old nodules were washed in 70% ethanol and rinsed in distilled water three times. Nodules were then fixed in formalin-alcohol (70%)-acetic acid (FAA) overnight and dehydrated in 70, 80, 90 and 100% ethanol for 30 min at each dilution. The nodules were then placed in a Samdri critical point dryer (Bozzola & Russell 1999). Nodule samples were Immersed in liquid CO2 at -10°C and exposed to a critical point pressure of 7.4 mPa. After the drying process, treated samples were mounted on stubs and sputter coated with gold-platinum for 60 sec. Images were observed on a Leo 453 scanning electron microscope operated at low beam voltage value (EHT) of 15 kv, and digitally recorded for further use. Ten replications of each leghemoglobin-containing nodules and ineffective nodules were prepared for observation. Statistical analysis were arranged in a randomized block design in the two greenhouse studies. Data were evaluated by 104 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 analysis of variance using the general linear models procedure of the Statistical Analysis Software (SAS Institute, Cary, NC). Significant differences {P < 0.05) between mean values of main effects were determined using the Ryan-Einot-Gabriel- Welsh Multiple Range Test separation procedure for all analyses. Results and Discussion Genetic characterization -ThQVQ are five described bean rhizobial species to this date: Rhizobium leguminosarum bv. phaseoli (Jordan), R. tropici (Martinez-Romero et al.), R. etii (Segovia et al. 1993), R, gallicum and R. giardinii (Amarger et al.). In addition, other isolates able to nodulate bean show distinct phylogenetic positions and may well represent other species (Eardly 1992). To establish the rhizobial species designation for the region under study, twenty-one isolates were genetically characterized on the basis of the polymorphism of 16S rRNA genes. The procedure differentiates between R. etli and R. tropici that are known to nodulate P, vulgaris L. in this continent (Segovia et al. 1993). Results indicated that the PCR-RFLP profiles of the 16S rRNA gene region obtained with nodule-producing strains detained during this study matched those of R. etli. In addition, the digestion patterns were not similar to those produced by R. tropici. Thus, the predominant populations of nitrogen fixing bacteria from this collection were phylogenetically assigned to the species R. etli. These results are in accordance to those of Segovia et al. (1993) who proposed that the species R. etli predominates in Mesoamerica, a region considered one of the main centers of origin of P. vulgaris. Rhizobial strain selection of the soil collection sites had a recent history of inoculation with N2-fixing bacteria with most soils being slightly alkaline. The predominant bean-nodulating popula¬ tion of R. etli in soils of the Lower Rio Grande Valley could be attributed to (a) introduction of the cultivation of beans by past agriculturalists who could have obtained seed from central or southern Mexico (Perez-Ramirez 1998), and (b) edaphic properties and favorable environmental conditions (c). Determinations of MATERON & EUBANKS 105 Table 1. Physical and biological characteristics where soil samples containing rhizobia able to nodulate Phaseolus vulgaris L, were collected from agricultural soils of the Lower Rio Grande Valley during the spring season of 2004. Isolate Soil series Coordinates Altitude pH (m) MPN rhizobia/ g soil UTPA-P5 Zapata loamy 26.3 L22N 98.41. 26W 146.86 7.53 2.1 X 10^ UTPA-P6 Reynosa silty clay 26.22.1 IN 98.47.44W 43.89 7.61 4.2 X 10^ UTPA-PO Ramadero loam 26.31.37N 98.41. 12W 13.41 7.65 8.3 X 10'^ UTPA-P17 Hidalgo sandy loam clay 26.28.57N 97.43.1 IW 3.05 7.84 4.0 X lO" UTPA-P18 Hargill sandy loam 26.26.43N 97.57.21W 13.10 8.01 1.1 X 10^ UTPA-P19 Wallacy fine sandy loam 26.28.55N 97.44.47W 14.93 7.75 8.3 X 10^ UTPA-P21 Wallacy fine sandy loam 26.28.56N 97.44.24W 13.41 7.72 4.1 X lO" UTPA-P34 Randado Cuevitas loamy 26.32.34N98.10.21W 22.55 7.81 5.4 X 10^ UTPA-P35 Evant silty clay 26.26.57N 98.34.05W 9.75 7.69 7.3 X 10^ UTPA-P39 Wallacy fine sandy loam 26.26.56N 98.07.07W 21.94 7.87 6.9 X 10^ UTPA-P44 Delfina fine loamy 26.26.54N 98.03.05W 21.33 7.84 8.7 X 10^ UTPA-P46 Delfina fine loamy 26.21.46N98.12.46W 25.91 7.73 5.5 X 10^ UTPA-P48 Delmita fine loamy 26.32.23N 98.09.13W 13.71 8.02 4.4 X 10'^ UTPA-P58 Delfina fine loamy 26.32.10N98.07.47W 17.98 7.81 5.1 X 10^ UTPA-P62 Harlingen clay 26.07.35N 97.49.24W 10.67 7.56 6.7 X 10^ UTPA-P66 Camargo silty loam 26.04.1 IN 97.48.54W 12.19 7.63 3.3 X 10^ UTPA-P69 Matamoros silty clay 24.04.20N 97.48.55W 8.23 7.60 2.1 X 10^ UTPA-P72 Raymonville clay loam 26.20.17N97.49.13W 27.43 7.45 6.4 X 10^ UTPA-P80 Hidalgo fine sandy loam 26.12.27N97.30.29W 12.19 7.63 5.9 X 10' UTPA-P82 Toscano clay 26.06.3 IN 97.48.36W 16.76 7.41 8.2 X 10^ bacterial numbers by the MPN method indicated that in most of the soils of the region the indigenous rhizobial population of bean rhizobia ranged between 1.1 x 10^ to 8.3 x 10"^ rhizobia per gram of soil (Table 1). This number is low but could be higher if stimulated by the presence of the host legume. Infection and structure of nodules preparations of nodule tissue were observed under the scanning electron microscope, it was evident that bacteroids from effective nodules were more visible and abundant inside peribacteroid membranes than those found in tissue preparations from ineffective nodules. Functional nitrogen-fixing bacteria were rod shaped and displayed a non- marked pleomorphism (Fig. 1). Conversely, membrane tissues of ineffective nodules had a low amount of bacteroids. These observations are in accordance to those of Floll & LaRue 1976, Bal 106 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Fig. 1. Examination of sections of nodule tissues of Phaseolns vulgaris L. by transmission electron microscopy. Image (a) shows an abundant number of bacteroids inside peribacteroid membranes of an effective nodule corresponding to strain UTPA-P66 at x 2,180. In contrast, emage (b) indicates a lack of bacteroids inside bacteroid membranes from a 1 0-day old ineffective nodule of strain UTPA-P69 at l,840x. MATERON & EUBANKS 107 et al. (1982) and Sprent & De Faria (1988). Infection threads were seen only in young cells of nodules. Bacteria were released from these cells soon after the end of the infection process. Infection threads appeared to have a defined sheath with a low number of bacteria visible inside. It was also evident that populations of the effective isolates were enclosed in large peribacteroid membranes (Fig. 1). Differences in the degree of adhesion between perio- bacteroid membranes and the envelope outer membranes may explain variations in the number of bacteriods enclosed by peribacteroid membranes in nodules of this host plant. Nodulation and N2-fixing potential. -SomQ researchers have reported that this legume species has a weakness in the ability to nodulate and fix nitrogen in symbiosis (Graham 1981; Graham & Halliday 1977; Graham et al. 2003). Contributing factors may include poor soil conditions under which the crop is grown (e.g., low soil organic matter, phosphorus content, intermittent or terminal drought - which are considered typical situations in the Rio Grande Valley region of Texas). Other factors that merit consideration are the short growing season of many cultivars, the limited energy supply to the nodule, and, in some areas, the effects of mineral nitrogen fertilization commonly applied to inoculated Phaseolus plants in the field (Graham & Halliday 1977; Michiels et al. 1998). Of the 83 sites examined for soil containing indigenous nitrogen-fixing bacteria for Phaseolus, only 20 of those sites contained rhizobia that were able to nodulate the host plant (Table 1). The rhizobial isolates varied markedly in their degree of nitrogen fixation as well as in size and shape of nodules produced. Several of the isolates formed nodules with an average dry mass tissue ranging from 160 to 620 mg per plant and were ranked as effective (E) or functional in fixing nitrogen (Table 2). Another cluster of strains were ranked as partially-effective (PE), and those producing lower nodule tissue with no visible traces of leghemoglobin were classified as ineffective (I) in fixing nitrogen. 108 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Table 2. Phaseolus vulgaris response to inoculation with isolates of R. etli under controlled growth conditions. Means with the same letter within a column are not significantly different at 5% probability level. Symbiotic effectiveness index calculated as shoot biomass of the mean of the inoculated plant divided by the mean of the shoot biomass of the non-inoculated control. E=effective, PE=partially-effective and I=ineffective nodulation (Beck et al. 1993). Strain 132 is a commercial strain from Becker Underwood (Ames, Iowa). Rhizobial isolate Shoot biomass (g) Total nodule mass (g) Number of crown nodule Number of lateral mass Symbiotic effectiveness index UTPA-P5 0.26“' 7.04'* 17.80“'* 7.26 (PE) UTPA-P6 1.96'*^ 0.18“' 8.17' 12.09'* 5.16(1) UTPA-PIO 3.25*'^''^ 0.38^'^'’ 17.52'“ 22.52' 8.55 (E) UTPA-P17 2 24def 0.17** 2.02* 13.00'*® 5.89 (I) UTPA-P18 3.03'“' 0.41“' 25.93“ 17.33“' 7.97 (PE) UTPA-P19 2.80'“' 0.46“' 11.00'“ 39.26“ 7.36 (PE) UTPA-P21 2 29“'f 0.28“' 9.90'“' 14.52“' 6.02 (I) UTPA-P34 2.73'“' 0.22“' 3.48'*® 18.74“' 7.18 (PE) UTPA-P35 2.47“'’’ 0.19“' 0.83®'' 18.57“' 6.50 (I) UTPA-P39 2. is" 0.20*®“ 1.38®“ 11.23*® 5.65 (I) UTPA-P44 2.78'“' 0.28“' 13.87'“ 8.77*® 7.31 (PE) UTPA-P46 2.23“'*' 0.16*^* 0.88*“ 15.00“'* 5.86 (I) UTPA-P48 3.10'“' 0.32'“ 8.28' 30.14“ 8.15 (PE) UTPA-P58 2 ^ycde 0.26'* 3.07'** 15.15“'* 7.81 (PE) UTPA-P62 2.83“' 0.20*® 4.17'* 13.04'* 7.44 (I) UTPA-P66 4.19'“ 0.55““ 13.24'“ 41.76“ 11.02 (E) UTPA-P69 1.95'*' 0.23“' 1.10®“ 22.30' 5.13 (I) UTPA-P72 1.64*s 0.18**“ 0.89*’’ 12.93'* 4.31 (I) UTPA-P80 4.29“ 0.55““ 40.30“ 18.1 1“'* 11.28 (E) UTPA-P82 3.78““' 0.62“ 43.08“ 27.50“' 9.95 (E) Strain 132 2 53“'f 0.27“' 6.63'* 21.5' 6.65 (I) UC+N 5.31“ 0* 0“ 0“ UC-N 0.38® 0* 0“ 0“ 1.00 MATERON & EUBANKS 109 Isolates producing ineffective nodulation were not associated with plants having higher amounts of shoot biomass and thus had the lowest symbiotic effectiveness indices (Table 2). Strains P82, P80 and P66 produced an average of 55.8 mg of nodule tissue, a significantly higher (P < 0.05) amount of tissue biomass than any of the other rhizobial strains tested. It has been known that crown nodulation is an indication of competitive nodulation success in the early events of root hair infection by rhizobia colonizing the rhizosphere (Graham & Halliday 1977; Graham 1981). All rhizobial isolates formed nodules in both the primary and secondary root system. In only 4 cases (strains PI 8, P44, P80 and P82), the highest proportion of nodules was located within the crown or upper section of the primary root. Strain P80 formed 40 effective functional nodules in the crown and 18 on the lateral root system. On the other hand, strains P19 and P66 produced a higher (P < 0.05) number of nodules in the lateral root system as compared to those produced by other rhizobial isolates (Table 2). Nodule distribution due to strains P80 and P82 was more concentrated at the crown section of the primary roots than in the secondary root system. Although P19 was not a good nitrogen fixer, it had 78% of its nodules positioned in the lateral root system. Despite a uniform cell density of 10^ cells applied to the emerging root, the response of strain P19 may suggest that early root hair infection events for nodule formation at the crown area could involve a complex series of physiological events. The commercial inoculant strain, Becker Underwood No. 132, currently used to inoculate crops of P. vulgaris in the region, proved to be inferior to most of the indigenous rhizobial isolates in their ability to fix nitrogen. Shoot and nodule tissue mass produced using this inoculant was significantly lower {P < 0.05) as compared to strains P66, P80 and P82 (Table 2). Furthermore, strains P80 and P66 produced a comparable biomass {P > 0.05) as that produced by plants that received no inoculation but were supplemented with 110 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 mineral nitrogen. The ability of these indigenous rhizobia to produce nodules with visible interior red pigmentation due to leghemoglobin, indicated that they were effective with a potential to establish successful symbiotic associations. The elite nitrogen¬ fixing strains identified in this study, were able to produce up to eleven times more biomass than plants with nitrogen deficiency symptoms. Conversely, the shoot biomass of plants that received no inoculation and no mineral nitrogen was lower {P < 0.05) than any shoot mass value of any rhizobial strain under study. No nodules were observed in plants of the uninoculated or mineral nitrogen control treatments. The potential of using strains P66, P80, and P82 as local inoculants for bean plants depends, however, on their growth, competitiveness and persistence under the soil conditions of the Lower Rio Grande Valley. Among these conditions, pH, drought, and microbial competitors should be evaluated. More information is required about the performance of these elite rhizobial isolates under field experimentation to further establish their value as commercial inoculants. These results point out the benefits which could be achieved by selecting efficient and competitive strains from populations of indigenous nitrogen-fixing bacteria from agricultural soils of the region. It was evident that these elite isolates are better adapted to local environmental conditions than the introduced commercial inoculant product. The latter resulted in low symbiotic effectiveness indices comparable to plants with ineffective nodulation (Table 2). Selecting highly effective rhizobia for inoculants should not neglect the ability of the introduced bacteria to compete with the indigenous populations of nitrogen-fixing bacteria. Introduction to agricultural fields, in different geographical areas, remains a decisive step when testing selected strains. These results point out the benefits which could be achieved by selecting efficient and competitive strains among natural populations of Rhizobium etli. MATERON & EUBANKS 111 Acknowledgements This work was partially supported by a University of Texas Faculty Research Council grant. We are indebted to Dr. Anxiu Kuang for allowing access to the Electron Microscopy Unit and sharing her knowledge in the preparation of samples for the scanning electron microscope. We are also grateful to Dr. Larry Zibilske, research soil scientist at the USD A Experiment Station in Weslaco (Texas), for allowing access to the environmentally-controlled plant growth facilities. Thanks to Drs. Frederick Zaidan and Robert Edwards for their critical review of this manuscript. Literatured Cited Aguilar, O. M., V. Lopez, P. M. Riccillo, R. A. Gonzalez, M. Pganano, F. H. Grasso, A. Puhler & G. Favelukes. 1998. Prevalence of Rhizobium e///-like allele in genes coding for 16S rRNA among the indigenous rhizobial populations found associated with wild beans from the Southern Andes in Argentina. Applied and Environ. Microbiol., 64(9):3520-3524. Anyango, B., K. L. Wilson, J. L. Beynon & K. E. Giller. 1995. Diversity of rhizobia nodulating Phaseolus vulgaris L. in two Kenyan soils with contrasting pHs. Applied Environ. Microbiol., 61(1 1):40 16-4021. Bal, A. K., S. Shamtharam & P. P. Wong. 1982. Nodulation of Phaseolus vulgaris L. by rhizobia two cross-inoculation groups. Appl. Environ. Microbiol., 44(4):965-97L Beck, D. P, L. A. Materon & F. Afandi. 1993. Practical Rhizobium-LQgumQ Technology Manual. International Center for Agricultural Research in the Dry Areas (ICARDA). Technical Manual No. 19, x+389 pp. Bozzola, J .J. & L. D. Russell. 1999. Preparation of Samples for the Scanning Electron Microscope. Electron Microscopy. Pp. 48-76 in Electron Microscopy, Principles and Techniques for Biologists. 2"^. Ed. Jones and Bartlett Publishers. Sudbury, MA., xxiii+670 pp. Eardly, B., L. A. Materon, N. H. Smith, D. A, Johnson, M. D. Rumbaugh & R. K. Selander. 1990. Genetic structure of natural populations of the nitrogen-fixing bacterium Rhizobium meliloti. Appl. Environ. Microbiol., 56(1):187-194. Eardly, B. D., J. P. W. Young & R. K. Selander. 1992. Phylogenetic position of Rhizobium sp. Strain Orl91, a symbiont of both Medicago sativa and Phaseolus vulgaris, based on partial sequences of the 16rRNA and nifH genes. Appl. Environ. Microbiol., 58(6):1809-1815. Graham, P. H., J. C. Rosas, C. Estevez, E. Peralta, B. Tlusty, J. Acosta-Gallegos & P.A. Arraes Pereira. 2003. Adressing edaphic constraints to bean production: the bean/cowpea CRSP project in perspective. Field Crops Res., 82(2-3): 179: 192. Graham, P. H. & J. Halliday. 1977, Inoculation and nitrogen fixation in the genus Phaseolus. Trop. Agric. Misc. Publ. 145. Univ. Hawaii Coll., 313-334 pp. Graham, P. H. 1981. Some problems in nodulation and symbiotic nitrogen fixation in Phaseolus vulgaris L. Field Crops Res., 4 (1):93-1 12. 112 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Roll, F.B.& T.A. LaRue. 1976. Genetics of legume plant host. Pp. 391-399 in Vol. II, Proceedings of the first international symposium on nitrogen fixation (W. E. Newton & C. J. Nyman, eds.). Washington State University Press, Pullman, WA. Materon, L. A. 1991. Symbiotic characteristics of Rhizobium meliloti in west Asian soils. Soil Biol. Biochem., 23(5):429-434. Materon, L. A. & J. Ryan. 1996. Effects of rhizobial inoculation and mineral nutrition of common Mediterranean pasture and forage legumes. Agric. Med., 126(l):64-74. Materon, L. A. & L. Zibilske. 2001. Delayed inoculation and competition of nitrogen¬ fixing strains in Medicago noeana (Boiss.) and Medicago polymorpha (L.). Appl. Soil Ecol., 17:175-181. Michiels, J., B. Dombrecht, N. Vermeiren, C. Xi & E. Luyten. 1998. Phaeolus vulgaris L. is a non-selective host for nodulation. FEMS Microbiol. Ecology, 26:193-205. Mrabet, M., R. Mhamdi, F. Tajini, R. Tiwari, M. Trabelsi & M. E. Aouani. 2005. Competitiveness and symbiotic effectiveness of a R. gallicum strain isolated from root nodules of Phaseolus vulgaris L. European J. Agron., 22(2):209-216. Mostasso, L., F. L. Mostasso, B. G. Diaz, M. A. T. Vargas & M. Hungria. 2002. Selection of bean {Phaseolus vulgaris L.) rhizobial strains for the Brazilian Cerrados. Field Crops Res., 73(1): 121-1 32. Perez-Ramirez, N.O., Rogel, M.A., Wang, E., Castellanos, J.Z. and Martinez-Romero, E., 1998. Seeds of Phaseolus vulgaris bean carry Rhizobium etli. FEMS Microbiol. Ecol. 26, pp. 289-296. Segovia, L., J. P. W. Young & E. Martinez-Romero. 1993. Reclassification of American Rhizobium leguminosarum biovar phaseoli type I strains as Rhizobium etli sp. nov. Intern. J. Systematic Bacteriol., 43:374-377. Smith, D. T. & J. L. Anciso. 2005. The Crop of Texas. Department of Soil and Crop Sciences. Texas A&M University. Extension Service. Technical Report SCS-2005- 01.63 pp. Sprent, J. I. & S. M. de Faria. 1988. Mechanisms of infection of plants by nitrogen-fixing organisms. Plant & Soil, 1 10(1):157-165. Tamimi, S. M. 2002. Genetic diversity and symbiotic effectiveness of rhizobia isolated from root nodules of common bean {Phaseolus vulgaris L.) grown in the soils of the Jordan Valley. Appl. Soil Ecology, 19(2): 183- 190. Thompson, J. D., T. J. Gibson, F. Plewniak, F. Jeanmougin & D. S. Higgins. 1997. The windows Clustal-X- interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res., 25:4876-4882. Turner, S. L. & J. P. W. Young. 2000. The glutamine synthetases of rhizobia: phylogenetics and evolutionary implications. Mol. Biol. EvoL, 17:309-319. United States Department of Agriculture (USDA). 2007. Natural Resources Conservation Service. Soil. Natural Cooperative Soil Service: http://websoilsurvey.nrcs.usda.gov/app/> Accessed 2007 Nov. 24. Vincent, J. M. 1970. A manual for the practical study of the root-nodule bacteria. International Biological Programme. Handbook No. 15, Blackwell Scientific Publications, Ltd., Oxford, UK. 164 pp. Zhang, A., S. Turner, X. Guo, H. Yang, F. Debelle, G. Yang, J. Denaire, J.P.W. Young & F. Li. 2000. The common nodulation genes of Astragalus sinicus rhizobia are conserved despite chromosomal diversity. Appl. Environ. Microbiol., 66(7):2988- 2995. LAM at: lmateron@utpa.edu TEXAS J. OF SCI. 60(2): 1 13-122 MAY, 2008 NOTABLE COUNTY RECORDS FOR ROLLING PLAINS ANGIOSPERMS OF NORTH CENTRAL TEXAS Allan D. Nelson and Jim R. Goetze Department of Biological Sciences, Box T-OlOO Tarleton State University, Stephenville, Texas 76401 and Science Department, Laredo Community College Laredo, Texas 78040 Abstract.-Based on data from fieldwork in Wichita County on a privately owned ranch located in the Rolling Plains region of Texas, thirty-nine species of flowering plants in 23 families are reported as new distribution and occurrence records for Wichita County, Texas. Twenty-one of these species represent major range extensions for the species in Texas. These new distribution and occurrence records provide baseline floristic data from a region that is poorly known compared to other regions of the state. Most of the new distributions are native but 31% are introduced species. Distribution data is critical for understanding native plant ranges and the spread of introduced species in the state. The flora of the Rolling Plains region of Texas is poorly known compared to other regions of the state (Turner et al. 2003a; 2003b). In fact, no comprehensive flora has been published for this region. Wichita County is located within the Rolling Plains region of Texas and little floristic work has been conducted in the county. The Rolling Plains form a partial boundary with the southeastern edge of the Panhandle region of Texas and extend southward to the northern Edwards Plateau and eastward to the West Cross Timbers regions of Texas. The Rolling Plains is considered a subsection of the Great Plains region of the central United States (Correll & Johnson 1970; Diggs et al. 1999). Rainfall ranges from approxi¬ mately 56cm (22 in.) in the west to almost 76cm (30 in.) in the eastern region with a summer dry period and corresponding high temperatures (Correll & Johnston 1970). Soils vary from coarse sands to heavy clays, or red-bed clays and shales with much of the area in crop or range land (Correll & Johnston 1970). The original prairie vegetation included tall and mid-grasses with short-grass species increasing under grazing. Mesquite {Prosopis glandulosa) is a common invader of all soil types, and heavy grazing by livestock (among other factors) has resulted in invasion of various 114 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Figure 1. Map showing the general loeation of floristic surveys within Wichita County. Inset shows location of Wichita County. species of grasses and forbs (Correll & Johnston 1970; Diggs et al. 1999). Study Area and Methods Plants were collected in Wichita County, Texas, mostly from a site 3 km N of Buffalo Creek Reservoir previously described by Stangl et al. (1992) as study site 2 - Goetze property or at Buffalo Creek Reservoir (Figure 1). Soils at the collecting sites were primarily clay loams. Much of the area is utilized for crop production and pasture lands. Plants were collected along roads, on the margins of wheat fields, in heavily grazed pasture lands, and around the margins of Buffalo Creek Reservoir. Collecting activities occurred in July 2003, April and May 2004, as well as May and September 2005. Plants were identified using Skinners & Mahler's Illustrated Flora of North Central Texas (Diggs et al. 1999) and the Manual of the Vascular Plants of Texas NELSON & GOETZE 115 (Correll & Johnston 1970). Nomenclature of native or naturalized plants was standardized and county records as well as range extensions are based upon the Atlas of the Vascular Plants of Texas (Turner et al. 2003a & b). Taxa are discussed alphabetically according to family. Voucher specimens are deposited in the Tarleton State University Herbarium (TAG). Results and Discussion Range extensions and reports of county records are critical in providing baseline floristic data with which to evaluate introduced species and better understand distribution of native species (Turner et al. 2003a). This study resulted in new distribution and occurrence records for 39 species from Wichita County in relation to information currently available in the Atlas of the Vascular Plants of Texas (Turner et al. 2003a; 2003b). Sixty-nine percent of the plants collected were native species. Eighteen species represent new county records for Wichita County but also have been reported (Turner et al. 2003a; 2003b) to occur in counties bordering Wichita County (Table 1). Twenty-one species represent major range extensions for plant species in Texas. While most of the flora collected in this investigation was native (69%), 31% of the major range extensions are introduced species. Each of these new species records is arranged by family and discussed individually within the accounts below. Family Apiaceae Torilis arvensis (Huds.) Link (Hedge-parsley; TAC 4007), an introduced species from the Mediterranean region known from disturbed habitats (Diggs et al. 1999), is chiefly found in the Blackland Praires and Edwards Plateau regions of Texas (Correll & Johnston 1970). The closest collections to the Rolling Plains locality in Wichita County are Grayson County to the east, Palo Pinto County to the southeast, and Garza County to the southwest (Turner et al. 2003a). 116 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Table 1. Floral records for Wichita County that have also been reported from bordering counties (Turner et al. 2003a & b) including Archer (A), Baylor (B), Clay (C), and Wilbarger (W). For species that have common names, they are included in parentheses at the end of the scientific name. Plants that are native (N) to north central Texas and introduced (I) are indicated. Accession numbers for the Tarleton State University Herbarium (TAC) are included. Family Species Bordering Counties N/I TAC Agavaceae Yucca arkansana Trel. (Arkansas yucca) A,C N 3992 Asteraceae Aphanostephus rarnosissimus DC. var. rarnosissimus (Plains lazy daisy) A, W N 4005 Helianthus maximiliani Schrad. (Maximilian sunflower) C N 4018 Senecio tampicanus DC. (Yellowtop) A N 4027 Sonchus asper (L.) Hill (Prickly sow-thistle) C I 4015 Boraginaceae Biiglossoides at^ensis (L.) l.M. Johnst. C 1 4021 Brassicaceae Lepidium oblongum Small (Veiny pepperweed) B N 4020 L. virginicum L. (Virginia pepper-grass) A N 4019 Cuscutaceae Cuscuta cuspidata Juss. ex Choisy (Cusp dodder) W N 4010 Nyctaginaceae Mirabilis linearis (Pursh) Heimerl (Linear-leaf four-o’clock) A,C N 4002 Onagraceae Oenothera laciniata Hill (Cut-leaf evening-primrose) C, W N 4026 Papaveraceae Argemone polyanthemos (Fedde) G.B. Ownbey C, W N 3991 Pedaliaceae Proboscidea louisianica (Mill.) Thell. (Common deviTs-claw) A N 4004 Poaceae Cynodon dactylon (L.) Pers. (Bermuda grass) C I 3996 Poa arachnifera Torr. (Texas blue grass) A N 4001 Scrophulariaceae Linaria texana Scheele (Texas toad-flax) A, C, W N 4023 Solanaceae Solanum rostratum Dunal (Buffalo-bur) A N 3989 Ulmaceae Celtis reticulata Torr. (Net-leaf hackberry) C N 4011 NELSON & GOETZE 117 Family Asteraceae Tragopogon dubius Scop. (Goat’s-beard; TAG 3997), introduced from Eurasia, is common in the west Cross Timbers and is migrating from the Panhandle to at least Grayson County in the east (Diggs et al. 1999). The collection in Wichita County helps affirm its migration from the Panhandle, where it is known from most of the counties of that region (Turner et al. 2003a). Family Brassicaceae Chorispora tenella (Pall.) DC. (Blue-mustard; TAC 4012), introduced from Asia, occurs in disturbed areas. This species has only recently been found in Texas (Diggs et al. 1999). The collection in Wichita County is the first report of this species from the Rolling Plains region of the state (Turner et al. 2003a). Family Cactaceae Echinocactus texensis Hoffer (Horsecrippler; TAC 3993) is native to Texas and usually found in sandy or limey soils in north central Texas, south and west to the South Texas Plains and Trans Pecos regions (Diggs et al. 1999). The closest collections to the locality in Wichita County are Foard County to the west and Young County to the south (Turner et al. 2003a). Mammillaria heyderi Muehlenpf (Nipple cactus; TAC 4016) is native to Texas and the closest collections to the locality in Wichita County are Randall County to the northwest and Brown County to the south (Turner et al. 2003a). The collection in Wichita County is the first report of this species from the Rolling Plains region of the state. Family Geraniaceae Geranium carolinianum L. (Carolina crane’s-bill; TAC 4006) is a native species occurring in woods, fields, and waste places in 118 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 various kinds of soils throughout most of Texas (Diggs et al. 1999). The collection from Wichita County extends its known range from Throckmorton County to the south and Montague County to the east into the northern Rolling Plains (Turner et al. 2003a). Family Moraceae Madura pomifera (Raf ) CK Schneid. (Osage-orange; TAC 4024) is found in stream bottoms, lower slopes, and waste places mainly in northeastern to north central Texas and southward into Central Texas and was originally native to only about 12 counties in this region (Diggs et al. 1999). Madura pomifera is most common in eastern Texas (Correll & Johnston 1970) but has been reported from localities in the Texas Panhandle (Turner et al. 2003a). The collection represents the first record of this species from the Rolling Plains. Family Phytolaccaceae Phytolacca americana L. (Pokeweed; TAC 3999) is a native species found in stream bottom woods and thickets or occasionally in disturbed areas throughout much of Texas (Diggs et al. 1999). The collection represents the first report of this species from the Rolling Plains (Turner et al. 2003a). Family Poaceae Andropogon glomeratus (Walter) Britton, Stems, & Poggenb. (Bushy bluestem; TAC 3998) is a native species known from roadsides and low moist areas throughout Texas (Diggs et al. 1999), but is more frequent in the eastern one-half of Texas than westward (Correll & Johnston 1970). The closest collections to the locality in Wichita County are Motley County to the west, Throckmorton County to the south, and Collin County to the southeast (Turner et al. 2003b). NELSON & GOETZE 119 Arundo donax L. (Giant reed; TAG 4009), introduced from the Mediterranean region, is known from nearly thoughout Texas (Diggs et al. 1999). The collection represents the first record of this species from the Rolling Plains outside Garza County to the southwest (Turner et al. 2003b). Avena fatua L. (Wild oats; TAC 4014), introduced from Mediterranean region, is scattered in Texas along roadsides and in other disturbed habitats (Diggs et al. 1999). The closest collections to the locality in Wichita County are Childress County to the west and Jack County to the southeast (Turner et al. 2003b). Bromus tectorum L. (Cheat grass brome; TAC 4003), introduced from Europe, is found in disturbed sites from north central Texas south and west to the Panhandle region of Texas (Diggs et al. 1999). This collection represents the first location of this species from the Rolling Plains of Texas (Turner et al. 2003b). Poa annua L. (Annual bluegrass; TAC 3990), introduced from Europe, is found in disturbed sites throughout Texas (Diggs et al. 1999). This collection represents the first location of this species from the Rolling Plains region of Texas (Turner et al. 2003b). Triticum aestivum L. (Wheat; TAC 3994) is a commonly cultivated species that was introduced from Asia. It frequently escapes into disturbed areas throughout Texas (Diggs et al. 1999). The closest collections to the locality in Wichita County are Randall County to the west and Young County to the south (Turner et al. 2003b). 120 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Family Polygonaceae Polygonum punctatum Elliott (Water smartweed; TAG 3995) is a native species occurring in wet areas throughout Texas, but it is more common in the eastern one-half of the state (Diggs et al. 1999). This collection represents the first location of this species from the Rolling Plains region of Texas. The nearest records are from Hemphill County in the Texas Panhandle and Tarrant County to the southeast (Turner et al. 2003a). Rumex crispus L. (Curly dock; TAC 4013), an introduced species from Europe occurring in disturbed, moist areas, is widespread throughout Texas (Diggs et al. 1999). The closest collections to the locality in Wichita County are Childress County to the west, Throckmorton County to the south, and Cooke County to the east (Turner et al. 2003a). Family Ranunculaceae Ranunculus sceleratus L. (Blister buttercup; TAC 4008) is a native species found in moist, often sandy soils in north central Texas, South Texas, the Edwards Plateau, and Rolling Plains (Diggs et al. 1999). This collection represents the first location of this species from the Rolling Plains (Turner et al. 2003a). Family Rosaceae Crataegus viridis L. (Greenhaw; TAC 4022) is a native species occurring in stream bottoms, fields, and slopes in east and southeast Texas, west to the Edwards Plateau and Rolling Plains (Diggs et al. 1999). This specimen represents the first collection of this species from the Rolling Plains region of Texas (Turner et al. 2003a). NELSON & GOETZE 121 Family Sapindaceae Sapindus saponaria L. (Western soapberry; TAC 4017) is a native species scattered throughout Texas. Sapindus saponaria occurs in stream bottoms, forest margins, and disturbed sites (Diggs et al. 1999). The closest collections to the locality in Wichita County are Childress and Cottle counties to the west. Young County to the south, and Grayson County to the east (Turner et al. 2003a). Family Scrophulariaceae Castilleja indivisa Engelm. (Entire-leaf paintbrush; TAC 4025) is a native species favoring sandy or silty soils within open woods, prairies, and disturbed areas in southeast and east Texas, west to the East Cross Timbers, but this species has been widely planted by the Texas Highway Department (Diggs et. al. 1999). This collection in Wichita County extends its range northward from Callahan County in the Rolling Plains (Turner et al. 2003a). Family Typhaceae Typha domingensis Pers. (Narrow-leaf cat-tail; TAC 4000) is a native species found in shallow water or wet ground throughout most of Texas (Diggs et al. 1999). The closest collections to this locality in Wichita County are Childress and Cottle counties to the west, Throckmorton County to the south, and Grayson County to the east (Turner et al. 2003b). Acknowledgments We would like to thank Edith and Oscar Goetze for allowing access to their property in Wichita County and for room and board while conducting fieldwork. 122 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Literature Cited Correll, D. S. & M. C. Johnston. 1970. Manual of the vascular plants of Texas. Texas Research Foundation. Renner, Texas, 1083 pp. Diggs, G. M., B. L. Lipscomb & R. J. O’Kennon. 1999. Shinners & Mahler’s Illustrated Flora of North Central Texas. Botanical Research Institute of Texas. Fort Worth, Texas, 1626 pp. Stangl, F. B., Jr., T. S. Schafer, J. R. Goetze & W. Pinchak. 1992. Opportunistic use of modified and disturbed habitat by the Texas kangaroo rat {Dipodomys elator). The Texas J. of Sci., 44(l):25-35. Turner, B. L., FI. Nichols, G. Denny & O. Doron. 2003a. Atlas of the Vascular Plants of Texas, Volume 1. Botanical Research Institute of Texas. Fort Worth, Texas, 648 pp. Turner, B. L., H. Nichols, G. Denny & O. Doron. 2003b. Atlas of the Vascular Plants of Texas, Volume IT Botanical Research Institute of Texas. Fort Worth, Texas, 240 pp. ADN at: nelson@tarleton.edu TEXAS J. OF SCI. 60(2):123-136 MAY, 2008 HERPETOFAUNAL INVENTORY OF CAMP MABRY, AUSTIN, TEXAS: COMMUNITY COMPOSITION IN AN URBAN LANDSCAPE Adam W. Ferguson*, Molly M. McDonough and Michael R. J. Forstner Department of Biology, Texas State University-San Marcos San Marcos, Texas 78666 "^Current address: Department of Biological Sciences Texas Tech University, Lubbock, Texas 79409 Abstract.-Amphibians and reptiles were surveyed using standard sampling techniques at Camp Mabry, Austin, Texas in 2004 and 2005 as part of the Texas National Guard’s biological inventory of the property, A total of 555 individuals representing 20 species were documented including six anurans, five turtles, four lizards, and five snakes. Amphibians compromised 55% of all captures followed by turtles with 36%. Of the 555 specimens of reptiles and amphibians captured, 77.4% consisted of the following five species: Bufo nebulifer, Syrrhophus marnocki, Trachemys scripta, Chelydra serpentina, and Rana catesbeiana. Results suggest that despite Camp Mabry’s potential habitat and its location within an ecoregion known to support a diverse assemblage of reptiles and amphibians, the herpetofaunal community is characterized by a lack of certain groups (e.g., large colubrid snakes and lizards) known from the surrounding area. Camp Mabry, in north central Austin, Travis County, Texas, is the headquarters for the Texas Military Forces. Despite its urban location, this 152-ha training site retains some native habitat, in contrast to the surrounding urban environment. It is this undeveloped acreage that makes Camp Mabry a unique site for reptiles and amphibians within the City of Austin. At this location, populations of native animals remain within a dense urban context (Ferguson et al. 2005; Matthews & Abbott 2005; McDonough et al. 2005). Camp Mabry is geographically within the border of the Edwards Plateau ecoregion (The Nature Conservancy 2004). The Edwards Plateau is well known for its unique flora and fauna, and supports a relatively high level of faunal endemics (Goetze 1998). Several endangered species, threatened species, and species of special concern have been reported from this region (TPWD 2003), including the Golden-cheeked Warbler (Dendroica chrysoparia). Barton Springs salamander (Eurycea sosurum), and Tobusch fishhook cactus (Ancistrocactus tohuschii). It is important that the 124 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 managers of Camp Mabry remain informed regarding the eurrent composition, relative abundance, and any subsequent changes in their faunal assemblages. To address this concern. Camp Mabry has authorized inventories to assess floral and faunal diversity on the base. Avian (Matthews & Abbott 2005), mammalian (McDonough et al. 2005) and insect (Abbott & Broglie 2006) inventories have been performed, however, until now a systematic survey of the herpetofauna had not been completed. This paper reports results for the baseline herpetofaunal survey of Camp Mabry, Austin, Texas. The design of this study allows for ease of replication and subsequent long-term monitoring of the herpetological assemblage on the facility. The main objectives were to (1) inventory the herpetofaunal community; (2) provide abundance, relative density, and distributional data on the species found; and (3) suggest any management options affecting these taxa on the facility that were seen as appropriate. Study Site Camp Mabry is located within the city limits of Austin, Texas. The site is bound to the north by 45^^ Street, the east by Loop 1 expressway, to the south by 35^^ Street, and to the west by residential properties (Figure 1). Eighty-one hectares are available for training activities and include numerous buildings, such as houses, offices, maintenance facilities, storage, and indoor training facilities (Wermund & Avakian 1994). However, a significant portion of the site (40%) retains a variety of native vegetation that has been detailed by Wermund & Avakian (1994). In 1996, The Nature Conservancy (TNC) defined four vegetation communities on Camp Mabry (Wolfe et al. 1996). These were meant to be descriptive designations within the site and do not strictly define divergent floristic community boundaries. The four designations provided by TNC were: Ashe Juniper-Oak Woodland (AJOW), Live Oak Woodland (LOW), Sugarberry-Elm FERGUSON, MCDONOUGH & FORSTNER 125 Figure 1. Map of Texas identifying Travis County and an aerial photograph of Camp Mabry illustrating its relation to the city of Austin and the Colorado River. Woodland (SEW), and Live Oak Savanna (LOS). In addition to the native plants, several invasive plant speeies were found to oeeur on the site (Reinecke et al. 2002). The most abundant of those inelude ehinaberry {Melia azedarach), tree-of-heaven (Ailanthus altissima), and Japanese ligustrom (Ligustrum japonicum). The locations of these plants were typically in or adjacent to disturbed sites 126 THE TEXAS JOURNAL OE SCIENCE-VOL. 60, NO. 2, 2008 including concrete waste piles, eonstruction disposal sites, and other development sites, which were seattered throughout Camp Mabry fragmenting vegetation eommunities (Wermund & Avakian 1994; Wolfe et al. 1996). Water resources, ineluding both ephemeral and permanent wetland areas are particularly important areas for reptile and amphibian fauna. There are several drainages on the northwest portion of the property that flow during rain events. The approximate bankful sizes of the two permanent ponds are 2.0 ha and 0.5 ha with depths of 3.7 meters and 2.1 meters, respeetively (Farquhar et al. 1996). Materials and Methods Vegetation communities (Wolfe et al. 1996) were subdivided into eleven distinct zones. AJOW was divided into WAl, WA2, and WA3. LOW was divided into Wl, W2, W3, W4, and W5. SEW was divided into RIPl and RIP2. Category eleven, manicured area (MA), encompassed all areas of the facility managed as manicured lawns or parkland and represented the LOS category of vegetation (Ferguson et al. 2005). These zonal descriptions allowed a systematic inventory of the property conducive to future monitoring designs, while emphasizing the current vegetative structure of the facility. Several survey techniques were used within each of the ten regions in an attempt to detect all possible reptiles and amphibians on the facility. Different species as well as ecological groups of reptiles and amphibians are often sampled more successfully using survey techniques speeific to particular taxa (Parris et al. 1999; Sutton et al. 1999; Christiansen & Vandewalle 2000; Jenkins et al. 2003; Ford & Hampton 2005). Therefore, targeted species survey techniques were employed to maximize ehances of documenting all reptile and amphibian taxa. For example, snakes and lizards were targeted by drift fences with funnels, but also detected using visual encounter searches and time constrained searching (Christiansen & Vandewalle 2000). Funnel placement was coneentrated along RIP 1 and RIP 2 where erosion control fences had been previously FERGUSON, MCDONOUGH & FORSTNER 127 installed, although opportunistic placement of other funnel traps did occur. Due to high levels of human activity and constant training activity on the base, pitfall traps with drift fence arrays were not used, although with proper planning and placement this technique could be applied in the future. Anurans were also monitored using standard call surveys (Heyer et al. 1994; Ford & Hampton 2005) at four designated sites (Ferguson et al. 2005). Call surveys were performed for 20-minute intervals after a five-minute acclimatization period and standard weather variables including relative humidity, temperature, and wind speed were monitored using a Kestrel hand-held weather meter. Abundances of anurans were estimated for large choruses (Heyer et al. 1994) by locating individuals calling along the shore with artificial lights to obtain relatively accurate numbers. To minimize double counting, anuran individuals were included in the results only if they were counted in different aggregates or locations during surveys. This helped to reduce over-estimating individuals at the four call sites and provided locations of other known anuran breeding sites on the property. Aquatic sampling with eel pots in the creek and springs found in W1 sought to document karst dwelling salamanders (e.g., Eurycea spp.). Turtles were trapped using hoop nets (Steen & Gibbs 2004) and basking traps as well as by visual surveys of basking individuals. All terrestrial and aquatic species encountered during visual surveys were recorded. The site was visited four times a month at all times of the day and night, totaling 60 visits averaging 9.2 person hours per visit, for a total of more than 700 person hours. Nocturnal road searches were conducted along a pre-determlned 7.7-km stretch of roads that encompassed all of the habitat types, including the savannah (McDonough et al. 2005). A minimum of two persons performed road searches, driving no faster than 8.0 km/h, with one individual operating a spotlight and the driver scan¬ ning the roadways for reptiles and amphibians. Road cruising was conducted at various times of the night across all months surveyed. 128 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 All species encountered were opportunistically collected by hand capture, rubber bands, lizard nooses, and nets (Anderson 1965). Taxonomic identification followed Dixon (2000). Each individual animal had standard measurements recorded, was marked, and released at their point of capture. Marking systems employed were approved by Texas State University’s International Animal Care and Use Committee (lACUC #03FF16A316_03) and the Society for the Study of Amphibians and Reptiles and included toe clippings for lizards and anurans, drilling of carapace marginal scutes on turtles, and ventral pre-anal scale clips for snakes (Femer 2007). Anurans were collectively marked based on capture year (e.g., toe four for 2004 and toe five for 2005) and all other individuals were assigned unique numbers via a combination of toe clippings. Most of the species encountered are considered common throughout their distribution and are well represented in vertebrate collections from surrounding areas. This fact, in conjunction with the low abundances of each species observed at the site resulted in the decision that the taking of traditional voucher specimens was not warranted. However, tissue samples were collected from each taxa and were accompanied by photographic vouchers when possible. All tissue samples were deposited in Dr. Michael R. J. Forstner’s frozen tissue collection housed in the Biology Department of Texas State University-San Marcos. Exact locations of captures were recorded using a Garmin GPS II Plus unit. Results During the 14-month study period (21 April 2004 - 7 May 2005), 555 animals were documented representing 20 species, including six anurans, five turtles, four lizards, and five snakes (Table 1). An additional species, a juvenile Texas rat snake {Elaphe obsoleta) was salvaged in October of 2005 during a bat survey of Camp Mabry and is deposited in the Angelo State Natural History Collection, Angelo State University, San Angelo, Texas (ASNHC #14243). Amphibians compromised 55% of all captures followed by turtles with 36%. Of the amphibians. Coastal Plain toad {Bufo nebulifer) Captured after the survey period FERGUSON, MCDONOUGH & FORSTNER 129 o ^ p K) — U» ^ Ul ON NJ 45- On K) OJ U> I—* oo 4^ (-/i to 4:^ ^ NO U> k) OO 45- 4^ tsj to 45-. ►— -O U> 4^ >— • Ik> OJ to u> oo 45^ Oi ^ 0> — OOON45- N04i.aN ONOilOOJ45-U> •o45-tOoo *— 45>OU>4:».nO JO — — OOst-n ppppo ^pOp pp_4^p_— ‘p L>N0 45.tO toC4;fc-.Lrtty»bo P O O ON p NO 4s». to to io Cj kj (ON Table 1. Amphibian and reptile species captured by method for Camp Mabry, Austin, Texas 21 April 2004 - 7 May 2005. Species Time Road Basking Hoop Minnow Calls Incidental Total Searches Searches Trap Net Trap 130 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 was the most abundant (ca. 165) while the red-eared slider (Trachemys scripta) was the most abundant reptile (ca. 80). Among the four species of lizards captured, the ground skink (Scincella lateralis) was the most frequently encountered with 18 individuals (56% of lizards captured) followed by the introduced Mediterranean gecko (Hemidactylus turcicus) with seven individuals or 22%. Among the five species of snakes captured, rough earth snake {Virginia striatula) was the most frequently captured with five individuals representing 29% of snake captures (Table 1). Turtle trapping resulted in the capture of 132 unique individuals including 69 T. scripta, 31 common snapping turtle (Chelydra serpentina), 24 Texas river coolers {Pseudemys texana), and two stinkpots {Sternothorus odoratus), all of which received unique marginal scute marks. Of the 555 specimens of reptiles and amphibians captured, 77.4% were five species (Table 1), Bufo nebulifer, cliff chirping frog {Syrrhophus marnocki), Trachemys scripta, Chelydra serpentina, and bullfrog {Rana catesbeiana). Additionally, 13 of 20 species were represented by five individuals or less including the cricket frog (Acris crepitans) and narrowmouth toad (Gastrophryne spp.), both heard only once. Of the survey methods, time-constrained searches and incidental encounters yielded the greatest species richness, with 15 and 11 species respectively. All five snake species and two lizards {Hemidactylus turcicus and Sceloporus undulatus) were captured during either time-constrained searches or incidental encounters. For both reptiles and amphibians, time-constrained searches yielded the highest capture rate among active collecting techniques (one capture per 3.2 hr). Nocturnal searches lead to the capture of only four individuals composed of two species, Bufo nebulifer and Chelydra serpentina, with a capture rate of one capture per 4.9 hr. Among indirect methods, hoop net trapping provided the highest capture success for turtles (one capture per 0.13 trap night) whereas FERGUSON, MCDONOUGH & FORSTNER 131 anuran call surveys provided the highest detection rates for anurans (one capture per 4,5 min). Discussion This survey provides strong evidence that Camp Mabry’s herpetofaunal community is characterized by a subset of reptiles and amphibians known to oceur in this region of Texas (Dixon 2000). The natural histories of the five dominate speeies, Bufo nebulifer, Syrrhophus marnocki, Trachemys scripta, Chelydra serpentina, and Rana catesbeiana best explains the presenee, and high relative abundance of these speeies on the site. With the exception of Syrrhophus marnocki, whose range is restrieted to the Edwards Plateau ecoregion of Texas, all of these speeies are found aeross a broad geographie range (Conant & Collins 1998). Such broad scale distributions inherently imply a level of tolerance in both habitat and food resourees in order to fill regionalized niehes and may help explain why this suite of five species continues to thrive in Camp Mabry’s urban context. Based on these survey efforts, it appears that the current herpetofaunal diversity of this Texas National Guard site is characterized by a subset of reptiles and amphibians and a lack of particular species (e.g., large colubrid snakes) when compared to herpetofaunal surveys of other Travis County sites and potential speeies known to oceur in the county. After removing species unlikely to oecur based on the availability of habitat, food resources, and other environmental factors, the estimated number of potential species oeeurring on Camp Mabry is seventy-eight (Dixon 2000). This study found only 25%, or 20, of these potential species during the surveys on Camp Mabry. Even after refining the potential species list there are a number of notable speeies that were not encountered during the survey. Although published works on herpetofaunal communities exist for Travis County and surrounding areas (e.g., Streeker & Williams 1927; Streeker 1930; Smith & Bueehner 1947), relatively few inventories of proximal properties have been conducted in recent 132 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 years. O’Connor (2003) conducted a herpetofaunal survey on the Lady Bird Johnson (LBJ) Wildflower Center in west-central Travis County and found 13 species of reptiles and amphibians. Although the total number of species documented was less than the 20 species recorded for Camp Mabry, the author did not survey for aquatic turtles. Turtles account for 25% of the species diversity at Camp Mabry. O’Connor (2003) documented a much higher diversity of snakes with seven total species including large species such as the bullsnake {Pituophis catenifer), coachwhip {Masticophis flagellum), and western diamondback rattlesnake {Crotalus atrox). Two noteworthy snakes that were located on the LBJ Wildflower Property but absent from Camp Mabry were the rough green snake (Opheodrys aestivus) and the black-necked garter snake (Thamnophis cyrtopsis), both of which are known to occur near Austin (Milstead 1953). A survey of The Nature Conservancy’s Barton Creek Habitat Preserve (TMM/TNHC 2008) highlights other typical species of the Edwards Plateau not encountered at Camp Mabry. To date, the survey of BCHP has documented 27 reptiles and amphibians including eight species of snakes and five species of lizards not found at Camp Mabry. Some common species detected at BHCP but not located at Camp Mabry include green anole {Anolis carolinensis), Texas spotted whiptail {Aspidoscelis gularis), and ribbon snake {Thamnophis proximus). Other species of reptiles and amphibians expected yet undetected on Camp Mabry include three species of caudates {Amby stoma texanum, Eurycea neotenes, and Plethodon albagula), five species of anurans {Bufo woodhousii, Hyla cinerea , Rana sphenocephala, R. berlandieri, and Scaphiopus couchi), and four turtle species {Kinosternon flavescens, K. subrubrum, Terrapene carolinensis, T. ornata). However, only one of these species {Rana sphenocephala) was detected during two surveys of other Travis County sites (O’Connor 2003; TMM/TNHC 2008). The reduced diversity of Camp Mabry’s herpetological community is also evident when comparing capture rates to other FERGUSON, MCDONOUGH & FORSTNER 133 inventories with similar intensity. A vertebrate inventory of Richland Creek Wildlife Management Area, Freestone and Navarro counties, Texas (Ryberg et al. 2004) found nocturnal road searches provided the highest capture rate (one capture per 3.5 min) totaling 141 individuals of 11 species, which differed from the results of this study, where only 4 individuals of two species were captured (one capture per 4.9 hr). Although Richland Creek WMA is much larger (ca. 5,583 ha) than Camp Mabry, efforts were similar among sites n = \1 fox 18.8 hr and « = 15 for 19.5 hr, respectively. Similarly, survey results of Camp Maxey, Lamar County, Texas, detected 44 species of herpetofauna representing 5009 individuals, or 54% of the species expected from known geographic ranges (Ford & Hampton 2005). Again, Camp Maxey is larger (ca. 2570 ha), and not bound by urban surroundings. The low number of individuals captured (ca. 555), low species richness, and dominance of a particular suite of reptiles and amphibians could possibly be the result of three factors: nonnative plants and animals, unsuitable habitat, and the urban context. Each of these factors plays a role in limiting diversity of reptiles and amphibians at Camp Mabry. One of the most substantial threats to Camp Mabry’s herpetofaunal diversity is the large populations of nonnative animals such as roof and Norwegian rats {Rattus spp.) and red imported fire ant {Solenopsis invicta) which are known to prey upon a variety of vertebrates (Clark 1982; Wojcik et al. 2001). The presence of the nonnative rats at Camp Mabry, especially at the high densities found by McDonough et al. (2005) could negatively affect the herpetofaunal community found on Camp Mabry. These nonnative rodents compete for space and often reside in habitats highly suited for native reptiles and amphibians including old logs, debris piles, under large rocks, and even in trees. The imported red fire ant provides nearly the same conflicts in several of these habitat situations as well. In addition to these non-native species, several house cats {Felis catus) were documented across Camp Mabry, 134 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 which have also been shown to impact native reptile and amphibian assemblages (Henderson 1992). Finally, habitat degradation, which results from both nonnative vegetation and anthropogenic disturbance, is a likely factor influencing diversity. Nonnative vegetation, which most often originates from anthropogenic sources, can cause serious alterations of a habitat and has been documented to cause declines in reptile and amphibian communities (Minton 1968; Gibbons et al. 2000). Camp Mabry’s tracts of chinaberry, tree of heaven, and Japanese ligustrum create relatively large areas of unsuitable habitat for native herpetofauna (Reinecke et al. 2002). Conclusions These survey results indicate that Camp Mabry’s herpetofaunal community is diminished in terms of potential diversity. Over 74% of total specimens captured were made up of five species: Bufo nebulifer, Syrrhophus marnocki, Trachemys scripta, Chelydra serpentina, and Rana catesbeiana. Of additional concern, is the lack of certain species groups, most notably large colubrid snakes and lizard species, which have been documented on neighboring properties in Travis County. The prevalence of both nonnative plants and animals, including Rattus spp. and fire ants, may be influencing the herpetofaunal community on Camp Mabry and adding to the limited diversity documented during this survey. This survey provides the first systematic inventory of Camp Mabry’s herpetofaunal community, establishing baseline data needed for monitoring the responses of Mabry’s reptiles and amphibians to active management practices already underway at Camp Mabry. Acknowledgements We would like to thank Dawn Johnson and the Texas Military Forces for supporting this project with both funding and on-site assistance. Gratitude is extended to all the security and other staff members of Camp Mabry. We would like to thank Travis LaDuc, Assistant Curator of Herpetology at the Texas Memorial Museum at The University of Texas at Austin, for conversations regarding their herpetofaunal survey of the Barton Creek Habitat Preserve in Travis FERGUSON, MCDONOUGH & FORSTNER 135 County, Texas. Additionally, we would like to thank Randy Simpson, Richard W. Manning, Kathleen O’Connor, Niara Pinto, Jake Jackson, Sue Reilly, Zach Gompert, and Lauren Lucas for assistance in the field. Literature Cited Abbott, J. C., & D. Broglie. 2006. Insect Survey Results for Camp Mabry, Travis County, Texas. Unpublished report. University of Texas, Austin, 55 pp. Anderson, R. M. 1965. Methods of collecting and preserving vertebrate animals. Nat. Mus. Canada Bull., Biol. Series no. 18., 199 pp. Conant, R., & J. T. Collins. 1998. A Field Guide to Reptiles and Amphibians of Eastern and Central North America. Third Edition, expanded. Houghton Mifflin Company, New York, 612 pp. Christiansen, J. L., & T. Vandewalle. 2000. Effectiveness of three trap types in drift fence surveys. Herpetol. Rev., 3 1(3): 158- 160. Clark, D. A. 1982. Foraging behavior of a vertebrate omnivore {Rattus rattus): meal structure, sampling, and diet breadth. Ecology, 63(3):763-772. Dixon, J. R. 2000. Amphibians and Reptiles of Texas. Texas A&M University Press, College Station, 421 pp. Farquhar, C. C., J. P. Maresh, P. Turner, M. Whiting, A. M. Price, S. Perry, B. Carr, D. Wolfe, C. & C. Liu. 1996. Biological Inventory of Texas Army National Guard Training Sites. Texas Parks and Wildlife Department, Resources Protection Division, Austin, 605 pp. Femer, J. W. 2007. A Review of Marking and Individual Recognition Techniques for Amphibians and Reptiles. Society for the Study of Amphibians and Reptiles, 72 pp. Ferguson, A. W., M. M. McDonough & M. R, J. Forstner. 2005. Herpetofaunal Diversity of the Texas National Guard’s Camp Mabry- Austin, Texas (May 2004 - May 2005). Unpublished report, Texas State University, San Marcos, 49 pp. Ford, N. B. & P. M. Hampton. 2005. The amphibians and reptiles of Camp Maxey, Lamar County, with comments on census methods. Texas J. Sci., 57(4):359-370. Gibbons, J. W., D. E. Scott, T. J. Ryan, K. A. Buhlmann, T. D. Tuberville, B. S. Metts, J. L. Greene, T. Mills, Y. Leiden, S. Poppy & C.T. Winne. 2000. The global decline of reptiles, Deja Vu amphibians. Bioscience, 50(8):653-666. Goetze, J. 1998. The mammals of the Edwards Plateau. Tex. Tech. Univ. Mus. Spec. Publ., 41:1-263. Henderson, R. W. 1992. Consequences of predator introductions and habitat destruction on amphibians and reptiles in the Post-Columbus West Indies. Carrib. J. Sci., 28 (1- 2):1-10. Heyer, R. W., M. A. Donnelly, R. W. McDiarmid & L-A. C. Hayek. 1994. Measuring and Monitoring Biological Diversity: Standard Methods for Amphibians. Smithsonian Institution Press, Washington, xviii+364 pp. Jenkins, C. L., K. McGarigal & L. R. Gamble. 2003. Comparative effectiveness of two trapping techniques for surveying the abundance and diversity of reptiles and amphibians along drift fence arrays. Herpetol. Rev., 34(l):39-42. Matthews, J .H. & J. C. Abbott. 2005. Bird Species Survey Report for Camp Mabry, Austin, Texas. Unpublished report. University of Texas, Austin, 28 pp. 136 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 McDonough, M. M., A. W. Ferguson & M. R. J. Forstner. 2005. Baseline Mammalian Survey of Camp Mabry Texas Army National Guard Training Site, Austin, Texas. Unpublished report, Texas State University, San Marcos, 41 pp. Milstead, W. W. 1953. Geographic variation in the garter snake, Thamnophis cyrtopsis. Texas J. Sci., 5(3):348-379. Minton, S. A., Jr. 1968. The fate of amphibians and reptiles in a suburban area. J. Herpetol., 2(3/4): 1 13-1 16. O'Connor, K. M. 2003. Herptile abundance and diversity in response to seasonal burning on a central Texas ashe juniper-live oak savannah. Unpublished M.Sc. thesis. Southwest Texas State Univ., San Marcos, 29 pp. Parris, K. M., T. W. Norton & R. B. Cunningham. 1999. A comparison of techniques for sampling amphibians in the forests of southeast Queensland, Australia. Herpetologica, 55(2):271-283. Reinecke, R., L. Clayton, K. Baird, R. Rolller & C. Knotts. 2002. Invasive Plant Species Survey, Camp Mabry, Texas, Unpublished report, Geo-Marine, Inc., Plano, 61 pp. Ryberg, W. A., L. A. Fitzgerald, R. L. Honeycutt, J. C. Cathey, & T. J. Hibbitts. 2004. Vertebrate inventory of Richland Creek Wildlife Management Area in eastern Texas. Southwestern Nat., 49(4):528-534. Smith, H. M., & H. K. Buechner. 1947, The influence of the Balcones Escarpment on the distribution of amphibians and reptiles in Texas. Bull. Chi. Acad. Sci., 8(1): 1-16. Steen, D. A., & J. P. Gibbs. 2004. Effects of roads on the structure of freshwater turtle populations. Conserv. Bio., 18(4):1 143-1 148. Strecker, J. K. 1930. A catalogue of the amphibians and reptiles of Travis County, Texas. Contrib. Baylor Univ. Mus., 23:1-16. Strecker J. K., & W. J. Williams. 1927. Herpetological records from the vicinity of San Marcos, Texas, with distributional data on the amphibians and reptiles of the Edwards Plateau region and central Texas. Contrib. Baylor Univ. Mus., 12:1-16. Sutton, P. E., H. R. Mushinsky & E. D. McCoy. 1999. Comparing the use of pitfall drift fences and cover boards for sampling the threatened sand skink {Neoseps reynoldsi). Herpetol. Rev., 30(3): 149-151. The Nature Conservancy. 2004. A Biodiversity and Conservation Assessment of the Edwards Plateau Ecoregion. Unpublished report, Edwards Plateau Ecoregional Planning Team, The Nature Conservancy, San Antonio, 121 pp. TMM/TNHC. 2008. Texas Memorial Musium/Texas Natural History Collections; University of Texas at Austin. Barton Creek Habitat Preserve website listing of Reptiles and Amphibians: Accessed July 2008. TPWD (Texas Parks and Wildlife Department). 2003. State of Texas Threatened and Endangered Species Listings. Texas Parks and Wildlife Department, Austin, 4 pp. Wermund, E. G. & A. J. Avakian. 1994. Physical Environment of Camp Mabry, Austin, Texas. Unpublished report, Texas Army National Guard, Austin, 37 pp. Wojcik, D. P., C. R. Allen, R. J. Brenner, E. A. Forys, D. P. Jouvenaz & R. S. Lutz. 2001. Red imported fire ants: impact on biodiversity. Am. Entomol., 47(1): 16-23. Wolfe, D. W., C. Liu, W. R. Carr, C. C. Farquhar, M. Goering, J. Maresh, P. A. Nguyen, S. Perry & G. Rowell. 1996. Land cover analysis of Texas Army National Guard Training sites. Unpublished report. The Nature Conservancy, Austin, 109 pp. AWF at: adamwferguson@gmail.com TEXAS J. OF SCIENCE 60(2): 137- 150 MAY, 2008 NOVEL AutoCAD METHOD FOR PERFORMING SURFACE ENERGETICS ANALYSES. Les N. McDaniel, Nicholas A. Romero, George P. Cobb and Gopal Coimbatore Department of Environmental Toxicology The Institute of Environmental and Human Health Texas Tech University, Box 41163, Lubbock, Texas 79409-1163 Abstract-This study reports the development of an economical apparatus and method for measuring contact angles of liquids on solid surfaces using a digital microscope and AutoCAD® 2005 drafting software. Digital images (JPG) of l.Opl droplets for a probe liquid triad (ultrapure water, diiodomethane, ethylene glycol) resting on the substrates of interest are produced. Using the AutoCAD® design window, images of droplets were overlaid with lines and shapes matching all profiles of the images. Once these reference lines were applied, contact angles (0) of the probe liquids at the three phase boundary were determined using AutoCAD® software and geometric principals. Multiple surfaces were analyzed using this method, including N-type silicon wafers, glass microscope slides, 130A thickness gold layers electrosprayed onto both N-type silicon wafers and glass slides, and a carboxyl-terminated self-assembled monolayer (SAM) deposited onto the gold layers. Ten replicates of each surface and treatments were tested independently to determine repeatability of the procedure. Contact angles determined for probe liquids were consistent and comparable to literature values. Surface free energy calculations for solids were calculated with the van Oss-Chaudhury-Good equation and were likewise consistent with literature values. This method of determining the contact angles of probe liquids on solid surfaces has proven reliable in confirming surface energy predictions based on the chemical and physical properties of those surfaces tested. Results support the premise that expensive optical apparatus and associated software are not required to perform accurate comparisons of surface wetting characteristics. Numerous fields of seience and engineering require detailed understanding of material surfaee characteristies in order to select or properly prepare surfaces. A common and familiar example is that paints are many times formulated for very specific applications. Spreading and adhesion of paint onto a target surface strongly influences the appropriateness of that paint for particular surfaces. For these reasons, paints intended for coverage of wood surfaces are often unsuitable for application to plastic or metal surfaces. Additionally, considerations must also be made concerning the surface characteristics of dried paints: Dried paints formulated for applications such as metal surfaces exposed to outdoor conditions 138 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 should have low surface energies in order to repel precipitation and thus protect the underlying materials from oxidation. In another entirely different and strongly contrasting utilization, surface wetting characteristics of plant and tree leaves have also been studied by environmental toxicologist to demonstrate exposures to automobile exhaust (Pal et al. 2002) and exposure to sulfur and deposition of heavy metals (Turunen et al. 1997). Under certain circumstances, observing changes in wetting properties of solid surfaces may be the only rapid and reliable method for confirming the presence or absence of modifications made to those surfaces. The application of self assembled mono- layers (SAMs) to gold-coated silica or glass is one such situation. A readily accessible and inexpensive technology capable of imaging these chemical monolayers is currently unavailable. Yet, by observing the contact angles of probe liquids of known polarities on these surfaces pre- and post-application of a SAM, researchers can compare changes in surface energies and determine the degree of success of the procedure. Hydrophobic/hydrophilic interactions between the solid surfaces and the applied probe liquids rapidly allow inferences to be made regarding any observed changes. Since surface energetics analyses and tensiometry are essential to many fields and have been in use for decades, many methods of acquiring and analyzing contact angles have been developed, and much debate has gone into determining which methods produce the most accurate and precise results. In fact. So many methods exist that to list and describe each here would not give fair treatment to any, and would be beyond the scope of any single manuscript. A review of literature comparisons reveals the Wilhelmy Balance method to be the benchmark, followed by the tilted plate and the sessile drop methods (Lander et al. 1993; Chibowski et al. 2002; Krishnan et al. 2005). While this need for testing the wetting properties of solid surfaces has widespread application, the associated costs may be MCDANIEL ET AL. 139 prohibitive to small laboratories where research grants may not be prevalent: Low-end commercial manual goniometers can cost $5,000-$6,000 new, and frequently require additional proprietary software and dedicated computer equipment in order to be fully functional. This report represents the use of inexpensive apparatus adapted for goniometric contact angle (0) analyses and an innovative use of AutoCAD‘S 2005 drafting software for determining 0 values. Materials and Methods Probe liquids and surface preparation -WmQty-mnQ percent pure diiodomethane (Alfa Aesar, Pelham, NH, USA), 99.0% ethylene glycol (VWR International), and ultrapure water were utilized as probe liquids. Selection of this triad was based on the simplification of surface free energy ) calculations via their utilization, and due to the preponderance of use in literature. Droplets of probe liquids (l.Opl) were applied to test surfaces via hand delivery with lO.Opl glass syringes (Hewlett Packard pn5 181-1267). Syringes were washed and autoclaved prior to use. Separate syringes were used for each probe liquid. The syringe for diiodomethane was wrapped in A1 foil during the procedure to prevent photodegradation of its contents. Droplet images were acquired within 3-5sec of deposition in order to minimize evaporative loss. Glass and silica wafers were immersed in chromic acid for 24 h, thoroughly rinsed with ultrapure water, and dried in a Precision Model 70D Laboratory 0ven for 12h at 100°C. Gold layers applied to glass slides and silica wafers were tested within one week of the electrospray gold deposition and atomic force microscopy. Self- assembled monolayers were likewise tested immediately upon completion (l-2h) of their deposition and drying. Self assembled monolayer A (C29H50O9S2; SensoPath Technologies, Bozeman, MT, USA) a dithiol-tethered SAM precursor molecule, was dissolved in an organic solvent and 140 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 deposited onto gold coated glass slides from solution. Upon completion, films were rinsed with anhydrous ethanol (Fisher Scientific, Pittsburg, PA 15219, USA) and dried under nitrogen stream. Apparatus -An adjustable instrument base with bubble leveler (Clay Adams, Parsippany, NJ 07054, USA) provided the foundation of this apparatus (Figure 1). Aluminum optical mounting dovetails (40 by 50mm) with !4”-20 counter bores (Thorlabs, Inc., 435 Route 206 North Newton, NJ 07860) were mounted onto the metal base as attachment points for the digital microscope and stage. A Digital Blue™ QX5™ Computer Microscope (e-bay, USA) was mounted by using two, /i” posts and 3” post holders (Thorlabs, Inc.) which were mounted to optical mounting dovetails. These mounting dovetails were joined to a single base-mounted dovetail via 40mm double dovetail clamps (Thorlabs, Inc.). This arrangement allowed horizontal and vertical adjustments of the microscope. To the second base-mounted dovetail, an in-house machined three-planar goniometer was similarly affixed. A 49.0cm aluminum plate was attached to the goniometer to serve as the stage. A 30.5cm section of 40mm optical dovetail rail was mounted to the metal base for attaching the diffuse light source. Analysis -Yifiy kilobyte, 512 x 384 digital images (JPG) of l.Opl droplets liquids resting on solid surfaces were acquired with a Digital Blue™ QX5™ Computer Microscope with the focus of the image on the profile of the droplet at its greatest height and diameter (Figure 2). Images were then imported into the AutoCAD® 2005 design window where tools were employed to overlay simple geometric shapes onto the images. First, a circle was overlaid onto the image and manipulated to match the profile of the droplet surface. This was accomplished by altering the diameter of the circle and moving the location of the mathematical center. Once this circle was best fit onto the image, a single straight line (Line 1) was applied to match the edge of the solid surface onto which the droplet is resting. The combination of this circle and Line 1 become the basis for which 0 calculations are performed. MCDANIEL ET AL. 141 Figure L Image of the apparatus developed for acquiring macro images of liquid droplets resting on solid surfaces (major components labeled). Digital Blue™ QX5™ Computer Microscope (A), stage (B), goniometer (C), diffuse light source (D), adjustable instrument base (E), Figure 2. Representative images of l.Opl droplets of test liquids resting on glass acquired with the apparatus developed during this study, (a) Diiodomethane, (b) Ethylene glycol, (c) MiliQ water. Contact angle was determined by the application of two additional straight lines and a rectangle to the image. Line 2 was formed by connecting the center of the circle and the point of intersection of the circle and Line 1, and was facilitated by the AutoCAD*^ design software: Points of intersection and center 142 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Figure 3. Droplet image demonstrating key elements of the AutoCAD analysis for determining theta. Lines A and B added to show true eenter and quadrants of the circle. points of circles are highlighted during modifieation of these files and are readily aeeessed by the user. The point where Line 2 intersects the eirele forms one vertex of a reetangle that has Line 2 as the bottom side. A reetangle was then inserted into the design window where the program auto-positions this shape so that two sides will parallel Line 1. The reetangle was rotated to a position where one side is eoincident with Line 2. A third line (Line 3) was then applied to the image using the side of the rectangle which lies closest to the eirele and is perpendieular to Line 2. At this juneture, all elements are in position for determining 0 for the droplet and solid surface (Figure 3). are ehosen from the AutoCAD® toolbar at the top of the design window. Two points within the design window are seleeted in order to reeeive output from this funetion. Two points along Line 3 are seleeted to generate 0. Theta is reported in degrees by the AutoCAD software in the Command dialog box at the bottom of the screen as “Angle in XY Plane =...”. MCDANIEL ET AL. 143 Table 1. Total surface tension and components (in mJ/m^) for water (W), diiodomethane (DIM), and ethylene glycol (EG) used for surface energy estimations. Liquid Yl 7l“ Yl' Yl' Water 72.8 21.8 51 25.5 25.5 DIM 50.8 50.8 0 0 0 EG 48 29 19 47 1.92 Surface free energy estimation.S\xxf?iCQ free energy estimations were ealeulated by inserting experimentally determined mean 0 values into the van Oss-Chaudhury-Good (vOCG) equation (l+cose)yL == 2(Vys^^VyL^^ + VysNyf + Vys'Vyp^) (Van Oss 2004). For the vOCG computational method, known reference values of the polar and dispersion components of a triad of pure liquids (Table 1) are utilized, along with experimentally determined 0 values for each. Three equations with three unknowns are then solved to obtain the values of ys^^, ys , and ys" . The total surface free energy is then calculated as ys^^^ = ys^^ + 2(Vys^ V ys' ). Microsoft® Excel® (XP, 2003) was used to generate surface free energy estimations. Single factor analyses of variances (ANOVAs) were performed on contact angles for each probe liquid between surface treatments (none, gold, SAM). Statistical comparisons were not made between data for glass and silica, nor were they made between the separate probe liquids. Results Mean contact angles for droplets of probe liquids resting on foundations of silica wafers and glass slides and the surface treatments were consistent, with standard errors generally increasing as the ys^^^ values increased (Table 2, Figures 4 & 5). Standard deviations for contact angles of droplets of all three probe liquids resting on each surface as determined by this method ranged from 1.07 to 9.59 (Table 2, Figures 4 & 5), with the poorer performance occurring for water on gold (glass foundation). 144 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Table 2. Sessile drop contact angles (in degrees) of probe liquids (Water, DilodoMethane, and Ethylene Glycol) on foundations of glass microscope slides and silica wafers with various surface treatments and total surface free energies (mJ/m^) calculated with the vOCG formula. Foundation Surface treatment 0 w 0DIM 0EG Glass None 32.4±1.07 50.9±2.96 19.0±2.94 46.00 Glass 20A gold 78.7±9.59 45.5±1.72 55.2±4.52 41.20 Glass SAM 37.9±9.21 22.5±3.31 21.2±6.83 62.84 Silica None 62.0±3.43 54.7±2.00 50.5±3.27 39.62 Silica 20A gold 60.1±4.36 37.6±3.10 42.3±3.56 50.91 Silica SAM 53.1±9.24 31.1±8.85 30.0±6.48 55.54 Surface free energy estimations for solid foundations and treatments, as expeeted, were highly variable depending on uppermost exposed layer contaeting the probe liquids, and ranged from 31.81 mJ/m^ (siliea, no surfaee treatment) to 62.84mJ/m^ (Au- eoated glass with SAM) (Table 2). One way ANOVA /^-values (j7=10, a=0.05) within probe liquids between surfaee treatments indieated extremely significant differences (/?<1.04E‘^) existed between all surfaee treatments for both glass slides and siliea wafers. Discussion The treated and untreated surfaees were expeeted to interact uniquely when brought into eontaet with the probe liquids. This was experimentally verified by eomparing the statistieal mean of the respective eontaet angles. Statistically significant and consistent probe liquid eontaet angle results between these surfaee treatments confirm the sueeessful deposition of gold and the subsequent deposition of the SAMs. The applieability of this method to researeh is demonstrated most fully by the eontaet angle results shown after the SAM deposition onto the gold surfaees. While the applieation of the gold was confirmed visually and through atomie foree mieroseopy prior MCDANIEL ET AL. 145 Figure 4. Contact angle means and standard errors for test liquids measured on glass microscope slides as determined with AutoCAD software. n=\0 for all six combinations of test liquids on solid surfaces. DIM=diiodomethane, EG=ethylene glycol, W=MiliQ water. Contact Angle Means (+/- SD) of Probe Liquids on Silica Foundation. 70 W DIM EG Probe Liquids. Figure 5. Contact angle means and standard errors for test liquids measured on N~ type silica wafers as determined with AutoCAD software. w=10 for all six combinations of test liquids on solid surfaces. DIM=diiodomethane, EG=ethylene glycol, W=MiliQ water. 146 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 to these analyses, no analytical method is capable of confirming the successful deposition of a chemical monolayer as rapidly and as contact angle determination. Although the standard deviations from the means of the 0 values were elevated above desirable levels for probe liquids applied to SAMs on both foundations, the statistical analyses demonstrate that relatively inexpensive equipment and software can be coupled in a manner which is capable of producing reliable results. The standard deviations were consistently elevated for contact angles of probe liquids on SAM’s and were randomly elevated for several other surfaces (Table 2). While no surface energetics analyses studies for a SAM identical or similar to the one used herein could be located, Lu et al. (1998) reported 0 values for the same triad of liquids on a hexadecyltrichlorosilane (HTS) SAM, similarly formed on both glass and silicon. Due to the molecular differences between these SAM’s, the contact angle means are not comparable, but the standard deviations for those measurements are useful and ranged from 0.8 to 5.0, whereas standard deviations in this study ranged from 3.31 to 9.24. Incomplete deposition of SAM’s onto the gold substrates (e.g., islands of exposed gold) is a likely explanation for elevating the standard deviations of observed means. Additionally, l.Opl droplets of probe liquids were approaching the lower limit of volume that can be accurately measured and hand delivered. This phenomenon was addressed when Lander et al. (1993) determined that the sessile drop method gave the lowest contact angles and was the least reproducible when compared to the Wilhelmy balance and tilted plate methods. In spite of these limitations, the sessile drop method continues to be a valuable tool while receiving ongoing scientific validation (Tadmor & Yadav 2007). Under ideal circumstances, probe liquid droplets should be mechanically measured and delivered, but the results show that hand delivery of droplets was a suitably reproducible. 0f all methods of contact angle determination currently in use, the nearest equivalent to the 0 values generated herein is the advancing contact angle (0a), regardless of which method is used. MCDANIEL ET AL. 147 although it has been concluded that needle presence in droplets affects the three phase boundary line (Lander et al. 1993). Since sessile droplets are at static hydraulic pressure, while advancing and receding contact angles (0r) are at increasing and decreasing hydraulic pressures, respectively, the sessile 0 for any given liquid will reside between 0a and 0r, but is more closely associated with 0a. Chibowski et al. (2002) reported that the 0a for water, DIM, and EG on glass microscope slides using the tilted plate method were 33.07 (±2.06), 46.12 (±0.74), and 24.95 (±2.36), which were in good agreement with corresponding experimentally determined 0 values of 32.4 (±1.07), 50.9 (±2.96), and 19.0 (±2.94) (Table 2). Chibowski et al. (2002) also found that 0a values for DIM and water on glass microscope slides generated with the syringe method were 47.75 (±1.24) and 29.91 (±1.69), respectively, which also strongly agree with corresponding experimental 0 values of 50.9 (±2.96) and 32.4 (± 1.07) (Table 2). Radelczuk et al. (2002) reported similar findings for water and EG on glass slides with 0a values of 33.6 (±1.1) and 48.8 (±0.8), respectively, which compared well with corresponding 0 values of 32.4 (±1.07) and 50.9 (±2.96) (Table 2). Contact angles determined on cleaned glass microscope slides and the subsequently calculated surface energies have become the predominant standard literature values for method and calculation comparisons. 0bserved estimate for cleaned glass slides (46.0mJ/m^) was in good agreement with literature values using the 0a values of the same triad of probe liquids (Radelczuk et al. 2002; ys^^^= 41.5mJ/m^, Chibowski et al. 2002; ys^^^= 43.5 ± 0.7mJ/m^ ). Surface free energy estimates for cleaned glass slides were also in strong agreement with the mean of ys^^^ calculations determined with 0 hysteresis for seven probe liquids (Radelczuk et al. 2002; ys =50.6 ±9.7mJ/m ) and through the 0a and 0r measurements of eight combinations of six probe liquid triads by the tilted plate method (Chibowski et al. 2002; ys^^^= 50.6 ±8.8mJ/m^ ). Since the ys estimates for cleaned glass slides were calculated with a commonly used triad of probe liquids and method, the agreement 148 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 seen between three findings and literature values demonstrate the suitability of this method of calculating 0. Volpe et al. (2003) stated that there is no “handbook-level” collection of contact angles of common liquids on common solids, nor measured or calculated surface free energies of common solids, and that 21^^ century scientists are not in agreement, even on the values of surface free energies of common solids. As such, it becomes apparent that variability between results remains inherent to the study of surface energetics, and that even large differences between data are to be expected. However, for the data presented herein, no such accommodations are required: Observed experi¬ mentally determined contact angles easily fall within range of frequently encountered data, while the calculated value for surface free energy of glass agrees strongly with values seen in current literature. Several steps of this procedure rely heavily on the patience and consistency of the analyst. The first and most important step is acquiring an analysis-appropriate image of the droplet on a solid surface. As mentioned earlier, it is essential that the focal point of the image be the profile of the droplet at its greatest height (true midpoint. Figure 3). This is to ensure that the points of contact between the droplet and solid surface are also clearly in focus. Macro photographs can be focused along a continuum of the droplet from leading edge to beyond. If the contact points at maximum droplet height are not properly in focus, application of the circle and Line 1 in AutoCAD® cannot be accurately achieved. It is essential to the success of the analysis that the droplet be in sharp focus thereby blurring the edge of the solid surface: Figures 2 and 3 show the blurring of the solid surface while the droplet and its reflection at point of contact remain clearly visible. Gaclawski & Urbaniak-Domagala (2007) emphasize the significance of capturing the reflection point clearly for the sessile drop method. For the analysis described, images should be of the highest quality resolution so pixilization is avoided: As images can be enlarged in the AutoCAD® design window during 0 determination to better fit MCDANIEL ET AL. 149 the components, increased resolution ensures accurately fitting the components necessary for analysis. An additional point is that the profile of a droplet resting on a surface is rarely the perfect arc of a circle bisected by a plane. This is especially true for droplets of low energy liquids laid onto low energy surfaces (eg., EG on glass, Figure 2b). While excessive droplet spreading may prevent the user from accurately applying a circle to such an image, the persistence of the user in fitting the circle to the best capabilities of the software increases the reliability and reproducibility of the results. Under instances where application of an ellipsoid to droplet profile is more appropriate, spheres in AutoCAD*® can be elongated along the horizontal axis accordingly without negatively effecting results. Surveys of existing literature for universal agreement on method of contact angle determination, acceptable probe liquids, and standard free energy values for common surfaces demonstrates that variability between these elemental components remains an open problem. Currently, investigators seem to have narrowed these problems and indications are that surface free energies are constant only under ideal conditions and, in reality, are best taken as the result of averaged values (Volpe et al. 2003, Chibowski et al. 2002). In spite of the limitations to the method described herein, developing an in-house apparatus and method for contact angle analysis is beneficial in many aspects. As demonstrated, the useMness of such a method in confirming successful surface treatments can be readily accomplished by novice operators. If these analyses are performed by using commonly accepted probe liquids, contact angle data can be inserted into surface free energy equations which result in estimations which concur strongly with literature values. In laboratories where surface energy studies play a small but significant role, such economical and reliable equipment and methods have great potential for applicability. 150 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 2, 2008 Acknowledgements Primary debt is to the United States Army’s Research, Development and Engineering Command (RDECOM) for providing the funding that made this project possible. Great appreciation also goes to Dr. Timothy Dallas, Ph.D. of Electrical Engineering at Texas Tech University for inspiring the development of this method and for his invaluable input and support. Literature Cited Chibowski, E., A. Ontiveros-Ortega & R. Perea-Carpio. 2002. On the interpretation of contact angle hysteresis. J. Adhesion Sci. Technol., 16(10):1367“1404. Goclawski, J. & W. Urbaniak-Domagala. 2007. The method of solid-liquid contact angle measurement using the images of sessile drops with shadows on substratum. MEMSTECH, 2007:135-140. Krishnan, A., Yi-Hsiu Liu, P. Cha, R. Woodward, D. Allara & E. A. Vogler. 2005. An evaluation of methods for contact angle measurement. Colloids and Surfaces B: Biointerfaces, 43:95-98. Lander, L. M., L. M. Siewierski, W. J. Brittain & E. A. Vogler. 1993. A systematic comparison of contact angle methods. Langmuir, 9:2237-39. Lu, Y., J. Drelich & J. D. Miller. 1998. Oleate adsorption at an apatite surface studied by ex-situ FTIR internal reflection spectroscopy. J. Colloid Interface Sci., 202(2):462-476. Pal, A., K. Kulshreshtha, K. J. Ahmad & H. M. Bell. 2002. Do leaf surface characteristics play a role in plant resistance to auto-exhaust pollution? Flora, 197(l):47-55. Radelczuk, H., L. Holysz & E. Chibowski. 2002. Comparison of the Lifshitz-van der Waals/acid-base and contact angle hysteresis approaches for determination of solid surface free energy. J. Adhesion Sci. Technol., 16(2): 1547- 1568. Tadmor, R. & P. S. Yadav. 2008. As-placed contact angles for sessile drops. J. Colloid Interface Sci., 317(l):241-246. Turunen, M., S. Huttunen, K. E. Percy, C. K. McLaughlin & J. Lamppu. 1997. Epicuticular wax of subarctic Scots pine needles: response to sulfur and heavy metals deposition. The New Phytologist, 135(3):501. van Oss, C. J. 2004. Use of the combined Lifshitz-van der Waals and Lewis acid- base approaches in determining the apolar and polar contributions to surface and interfacial tensions and free energies. J. Adhesion Sci. Technol., 16(6)669-677. Volpe, C. D., D. Maniglio, S. Siboni & M. Morra. 2003. Recent theoretical and experimental advancements in the application of van Oss-Chaudury-Good acid- base theory to the analysis of polymer surfaces I: General Aspects. J. Adhesion Sci. Technol., 17(1 1):1477-1505. LM at: les.mcdaniel@tiehh.ttu.edu TEXAS J. SCL 60(2), MAY, 2008 151 GENERAL NOTES REPRODUCTIVE CYCLE OF THE COMMON BARKING GECKO, PTENOPUS GARRULUS (SQUAMATA: GEKKONIDAE) FROM SOUTHERN AFRICA Stephen R. Goldberg Department of Biology, Whittier College Whittier, California 90608 The common barking gecko, Ptenopus garrulus occurs in western arid regions of southern Africa and ranges from little Karoo to Kruger National Park (Branch 1998). Hibbitts et al. (2005) studied the reproductive ecology of P. garrulus and Girard (1997) described mating behavior in captive specimens. Clutch sizes are in Haacke (1975), Pianka (1986), Pianka & Huey (1978). The purpose of this paper is to provide additional information on the reproductive biology of P. garrulus from a histological examination of museum specimens as part of an ongoing effort to characterize the reproductive patterns of geckos from southern Africa. Materials and Methods A total of 171 P. garrulus including 91 females (mean snout-vent length, SVL = 46.7 mm ±3.0 SD, range = 39-53 mm), 61 male (mean SVL = 47.0 mm ±3.6 SD, range = 38-58 mm), 10 subadults (mean SVL = 34.1 mm ± 1.6 SD, range = 33-37 mm) and 9 presumed neonates (mean SVL = 24.0 mm ± 1.5 SD, range = 21-25 mm) P. garrulus were examined from the herpetology collection of the Natural History Museum of Los Angeles County (LACM), Los Angeles, CA. Lizards were collected 1969, 1970, 1972 and 1976. For histological examination, the left testis and epididymis were removed from males and the right ovary was removed from females. Enlarged follicles (> 4 mm length) or oviductal eggs were counted. Tissues were embedded in paraffin and cut into sections of 5 pm. Slides were stained with Harris hematoxylin followed by eosin counterstain. Slides of testes were examined to determine the stage of the spermatogenic cycle. Slides of ovaries were examined for the presence of yolk deposition or corpora lutea. Statistical analyses were 152 THE TEXAS JOURNAL OF SCIENCE-VOL. 69, NO. 2, 2008 performed using Instat (vers. 3.0b, Graphpad Software, San Diego, CA). An unpaired ^test was used to compare P. garrulus male and female mean body sizes (SVL). Ptenopus garrulus from southern Africa examined from the herpetology collection of the Natural History Museum of Los Angeles County, (LACM), Los Angeles, California. BOTSWANA Kgalagadi District, 11 km S Tsabong (26°08’S, 22°28’E) LACM 83270, 83275, 83281, 83288, 83292, 83293, 83295, 83296, 83299, 83313, 83316, 83321, 83324, 83325, 83330, 83334, 83335, 83342, 83346, 83350, 83351, 83353, 83354, 83358, 83362, 83366, 83371- 83373, 83383, 83384, 83388, 83393, 83395, 83396, 83405, 83408, 83409, 83413, 83415, 83416, 83421, 83423, 83425, 83429, 83436, 83492, 83503, 83506, 83507, 83509, 83510, 83514, 83515, 83517, 83518, 83525; 9 km N, 11 km E Twee Rivieren (26°23’S, 20°43’E) LACM 83261,83263,83266. NAMIBIA Erongo Region, Walvis Bay (22°95’S, 14°50’E) LACM 127468, 127469; Karas Region, 89 km ENE Koes (26°00’S, 19°15’E) LACM 77292, 77293, 77299, 77300, 77302- 77307, 77310-77312, 77315, 77318-77323; Karas Region, 25 km WNW Helmering-hausen (25°88’S, 16°81’E) LACM 77047, 77050, 77052; Karas Region, 46 km N, 17 km E Aroab (26°22’S, 19°49’E) LACM 83206, 83207, 83210, 83211,83213,83215. REPUBLIC OF SOUTH AFRICA Northern Cape Province, 31 km N,100 km E Upington (28°13’S, 22H6’E) LACM 83138, 83146, 83148, 83150-83152; 24 km N, 83 km E Upington (28°17’S, 22°05’E) LACM 83223, 120 km N, 54 km W Upington (27°22’S, 20°43’E) LACM 83038, 83041, 83044-83051; 121 km N, 16 km E Upington (27°22’S, 2U25’E) LACM 83238, 83243, 83247; 129 km N, 65 km W Upington (27H7’S, 2r54’E) LACM 83055, 83059, 83064, 83066, 83072, 83074, 83076, 83077, 83080-83087, 83093, 83095, 83097, 83102, 83103, 83105-83111, 83113- 83115, 83117, 83120-83122, 83125-83129, 83131, 83132, 83134, 83136; Kalahari-Gemsbok National Park, 1 km W. Kameelsleep (25°45’S, 20°44’E) LACM 83156, 83160, 83163, TEXAS J. SCI. 60(2), MAY, 2008 153 83165, 83167, 83177, 83193, 83195, 83201-83204; 18 km S, 22 km E Witkoms (27°58’S, 21°32’E) LACM 83251, 83254; 50 km W, 7 km S of Vanzylsms (27°04’S, 21°48’E) LACM 83256, 83257. Results Males followed a seasonal testicular cycle (Table 1) in which three stages were represented: (1) Regression (non-reproductive) in which the germinal epithelium is exhausted and the predominant cells are Sertoli cells and spermatogonia; (2) Recrudescence (recovery) characterized by renewal of the germinal epithelium for the next period of sperm formation; primary and secondary spermatocytes are the predominant cells; (3) Spermiogenesis (sperm production) in which the seminiferous tubules are lined by clusters of spermatozoa and metamorphosing spermatids; the epididymides are packed with sperm. The peak period of sperm production occurred in austral spring when 100% of 31 males were undergoing spermiogenesis (Table 1). Recovery for the next period of sperm formation occurred during summer-autumn when all three stages (= regression, recrudescence, spermiogenesis) were present in the population (Table 1). Recovery is completed by the next spring. The smallest reproductively active male (spermiogenesis in progress) measured 38 mm SVL (LACM 83125) and was collected in October. There was no significant size difference (mean SVL) between males and females (unpaired t test). Monthly changes in the ovarian cycle are in Table 2. Egg production was limited to the right ovary. Females with enlarged ovarian follicles (> 4 mm), oviductal eggs or corpora lutea were recorded from October, December and January. Two females from April were undergoing early yolk deposition (marked by the presence of vitellogenic granules). There was no evidence (corpora lutea and early yolk deposition) in the same female to suggest more than one egg clutch is produced in the same reproductive season. Mean clutch size {n = 21) was 1.0 ± 0.0. The smallest reproductively active females measured 43 mm, follicles > 4 mm, (LACM 77323 from October and LACM 83223 from January) or, oviductal eggs, (LACM 77299 and LACM 77332 from October). 154 THE TEXAS JOURNAL OF SCIENCE-VOL. 69, NO. 2, 2008 Table 1 . Monthly changes in the testicular cycle of Ptenopus garrulus from southern Africa. Month n Regression Recrudescence Spermiogenesis September 9 0 0 9 October 20 0 0 20 November 2 0 0 2 January 6 4 0 2 February 6 3 2 1 April 15 2 6 7 May 3 1 1 1 Table 2. Monthly changes in the ovarian cycle of Ptenopus garrulus from southern Africa. Month n No yolk deposition Early yolk deposition Enlarged follicles > 4 mm Oviductal eggs Corpora lutea only Oct. 21 2 3 9 6 1 Dec. 5 4 0 0 0 1 Jan. 23 21 0 2 0 0 Feb. 25 25 0 0 0 0 Mar. 4 4 0 0 0 0 Apr. 12 10 2 0 0 0 May 1 1 0 0 0 0 Discussion In a previous study on the reproductive ecology of P. garrulus Hibbitts et al. (2005) reported peak reproductive activity for both sexes occurred in September and October. Data from this study as well as values in Pianka (1986) and Hibbitts et al. (2005) indicate that clutch size in P. garrulus consists of one egg. Hibbitts et al. (2005) reported that based on the presence of two eggs in different stages of development in 4 (8%) females, two egg clutches in the same year were possible; only the right ovary produced eggs. However, in contrast, no females were observed that would likely have produced more than one egg clutch in the same year (females with oviductal eggs and concomitant yolk deposition and/or females with corpora lutea and concomitant yolk deposition). This finding may be due to the small sample sizes. Hibbitts et al. (2005) reported reproductive activity commenced at smaller SVLs (male ==36 mm; female = 31 mm) than observed in this study. Gonads from ten P. garrulus from October in this study, (mean SVL = 34.1 mm ± SD, range = 33-37 mm were “extremely small” and TEXAS J. SCI. 60(2), MAY, 2008 155 consequently classified these lizards as sub-adults. These differences in onset of reproductive activity may suggest regional differences in the sizes that P. garrulus populations commence reproduction. It is not known if the two April P. garrulus that had initiated vitellogenesis (Table 2) would have deposited eggs or undergone follicular atresia with the yolk being reabsorbed (see Moodley & van Wyk 2007). Follicular atresia is common late in the reproductive season when follicles that do not complete vitellogenesis deteriorate (Goldberg 1973, Moodley & van Wyk 2007). Presumed neonates were collected in January {n = 6), February {n = 1) and May {n = 2). This is consistent with the observation of Haacke (1975) that young of P. garrulus hatch from early summer into autumn and Auerbach (1987) that eggs of P. garrulus probably hatch in late summer to autumn and possibly as early as October. To date reproductive cycles of African gekkonid lizards appear to fall into two types: (1) a seasonal cycle in which reproduction occurs mainly in spring-summer as found in P. garrulus described herein and in Hibbitts et al. (2005), Chondrodactylus angulifer, (Goldberg 2006a), Pachydactylus capensis, (Goldberg 2006b), Colopus wahlbergii, (Goldberg 2006c; Whiting et al. 2007), Pachydactylus bibronii, (Flemming & Bates 1995), Hemidactylus mabouia (Moodley & van Wyk 2007) and; (2) extended reproduction occurring through much of the year as appears to be typical for Lygodactylus, (Simbotwe 1983, Vitt 1986, Goldberg 2007, Goldberg 2008, Vences et al. 2004). Subsquent studies on additional species of geckos are needed before the variations in the timing of gekkonid reproductive cycles from southern Africa can be known. Acknowledgments I thank Christine Thacker (LACM) for permission to examine P. garrulus and Sean Kark (Whittier College) for assistance with histology. Literature Cited Auerbach, R. D. 1987. The Amphibians and Reptiles of Botswana. Mokwepa Consultants, Gaborone Printing Works, Botswana, 295 pp. 156 THE TEXAS JOURNAL OF SCIENCE-VOL. 69, NO. 2, 2008 Branch, B. 1998. Field Guide to Snakes and other Reptiles of Southern Africa, 3""* edn. Ralph Curtis Books Publishing, Sanibel Island, Florida, 399 pp. Flemming, A. F., & M. F. Bates. 1995. Male and female reproductive cycles of Bibron’s gecko Pachydactylus bibronii (Sauria: Gekkonidae) in the Free State province of South Africa. J. Afr. Zook, 109:409-422. Girard, F. 1997. Ptenopus garrulus maculatus Common barking gecko. Courtship and mating. Afr. Herp. News, 26:24-25. Goldberg, S. R. 1973. Ovarian cycle of the western fence lizard, Sceloporus occidentalis. Herpetologica, 29:284-289. Goldberg, S. R. 2006a. Reproductive cycle of the Namib giant ground gecko, Chondrodactylus angulifer (Squamata: Gekkonidae). Afr. Zook, 41 :308-3 1 1 . Goldberg, S. R. 2006b. Notes on the reproductive biology of the Cape Thick-toed gecko, Pachydactylus capensis (Squamata; Gekkonidae) from southern Africa. Bulk Chicago Herpetok Soc., 41:105-106. Goldberg, S. R. 2006c. Note on reproduction of Wahlberg’s Kalahari gecko, Colopus wahlbergii (Squamata: Gekkonidae) from southern Africa. Bulk Maryland Herpetok Soc., 42:140-143. Goldberg, S. R. 2007. Note on reproduction of the Cape Dwarf gecko, Lygodactylus capensis (Squamata: Gekkonidae) from southern Africa. Bulk Chicago Herpetok Soc., 42:195-196. Goldberg, S. R. 2008. Note on reproduction of the Painted Dwarf gecko, Lygodactylus picturatus (Squamata: Gekkonidae) from Kenya and Uganda. Bulk Maryland Herpetok Soc., 44:18-21. Haacke, W. D. 1975. The burrowing geckos of southern Africa, 1 (Reptilia: Gekkonidae). Ann. Transvaal Mus., 29:197-243. Hibbitts, T. J., E. R. Pianka, R. B. Huey & M. J. Whiting. 2005. Ecology of the common barking gecko {Ptenopus garrulus) in southern Africa. J. Herpetok, 39:509-515. Moodley, G. K., & J. H. van Wyk. 2007. Folliculogenesis and ovarian histology of the oviparous gecko, Hemidactylus mabouia (Sauria: Gekkonidae). Afr. J. Herpetok, 56:115-135. Pianka, E. R. 1986. Ecology and Natural History of Desert Lizards, Analyses of the Ecological Niche and Community Structure. Princeton University Press, Princeton, New Jersey, 208 pp. Pianka, E. R., & R. B. Huey. 1978. Comparative ecology, resource utilization and niche segregation among gekkonid lizards in the southern Kalahari. Copeia, 1978:69 1-701. Simbotwe, M. P. 1983. Comparative ecology of diurnal geckos {Lygodactylus) in Kafue flats, Zambia. Afr. J. Ecol., 21:143-153. Vences, M., P. Galan, K. Miramontes, & D. R. Vieites. 2004. Weak expression of reproductive seasonality in a dwarf gecko {Lygodactylus verticil latus) from arid south-western Madagascar. J. Arid. Environ., 56:329-338. Vitt, L. J. 1986. Reproductive tactics of sympatric gekkonid lizards with a comment on the evolutionary and ecological consequences of invariant clutch size. Copeia, 1986:773-786. Whiting, M. J., L. T. Reaney, & J. S. Keogh. 2007. Ecology of Wahlberg’s velvet gecko, Homolophis wahlbergii, in southern Africa. African Zook, 42:38-44. SRG at: sgoldberg@whittier.edu TEXAS J. SCI. 60(2), MAY, 2008 157 SITE FACTORS AFFECTING PINYON- JUNIPER OCCURRENCE IN THE CAPULIN VOLCANO FIELD OF NORTHEASTERN NEW MEXICO W. David Hacker and Jennifer L. Grosso Forestry Discipline, New Mexico Highlands University Las Vegas, New Mexico 87701 The Capulin Volcano Field covers approximately 207,192 km^ in Union and Colfax counties in northeastern New Mexico and southernmost Las Animas county in Colorado. The geology of the area is characterized by Pleistocene and Holocene extrusive volcanic activity through sandstone, shale and limestone of the Late Cretaceous Period (Trauger & Kelly 1987). Average annual precipitation and temperature for the area is 45.2 cm. and 9.6°C respectively (McCurdy 1994). Although, much work has been published on pinyon (Pinus spp.) - juniper {Juniperus spp.), little data have been reported on piny on-juniper stands from the Capulin Volcano Field. Species studied in this investigation include pinyon {Pinus edulis Voss.), one-seed juniper {Juniperus monosperma Sarg.) and Rocky Mountain juniper {J. scopulorum Lemmon.) [SAF Cover Type 239 (Larson 1980)]. Although not timber producing, these stands supply posts, fuelwood, and pinyon nuts. They provide nesting platforms for birds and edge and cover for wildlife and livestock. The objectives of this study were to locate piny on-juniper stands in the Capulin Volcano Field and determine what environmental factors are affecting their occurrence. Piny on-juniper stands were located and sampled on privately owned rangeland. Individual stands tended to be small (<3 ha) and surrounded by typical western Great Plains short grass dominated by blue grama {Bouteloua gracilis [Lag.]) and western wheatgrass {Agropyron smithii [Rybd.]). Data were obtained by approximating 158 THE TEXAS JOURNAL OF SCIENCE-VOL. 69, NO. 2, 2008 the center of each stand and randomly selecting the direction of a transect. Randomness was assured by following the direction of a second hand on a watch. Azimuth of the facing slope was recorded as was Universal Transverse Mercator (UTM) coordinates of the initiating point. Soil data were collected along transects at zero meters (the initiation point) and every five meters thereafter until the transect extended at least 10 meters beyond the edge of the stand. Each sample point was monumented by pin flags and numbered. Soil depth was measured with a soil probe. Soil depth was recorded by transect and flag number. Depth was measure to the nearest centimeter (cm) to 25 cm. Depths that exceeded 25 cm were recorded as >25 cm. Soil samples were collected at each sample point, recorded and returned to the laboratory. Mechanical analysis of soil samples was conducted using the method prescribed by Bouyoucous (1951; 1962). Soil pH was characterized using a glass electrode pH meter. Aspect was determined with the aid of a hand compass and azimuth was recorded. Percent slope was measured with a clinometer. Summaries of these data are shown in Table 1. Environmental variables were evaluated by means of stepwise regression with the presence of pinyon-juniper stands as the dependent variable. Environmental factors used as independent variables included: soil depth to 25 cm, percent clay, percent silt, percent sand, soil pH and percent slope. Data were standardized before statistical analysis was performed (SPSS 2003). Soil depth had the greatest impact on pinyon-juniper growth. Soil depth accounted for 31.2 percent of the variation of whether pinyon-juniper stands occurred and was negatively correlated. Soil pH accounted for an additional 7.8 percent of variation and was also negatively correlated with pinyon-juniper occurrence. Azimuth explained 4.2 percent of variation and was positively correlated. The resulting equation accounted for 43.2 percent of variation in seedling growth (P = 0.00). TEXAS J. SCI. 60(2), MAY, 2008 159 Table 1. Summaries of environmental data Variable Mean Standard Deviation Range Soil Depth (cm) 13.61 1.02 1-25 Soil pH 7.95 0.06 6.65-8.81 % Sand 39.09 10.83 12.00-72.72 % Silt 44.21 0.81 30.16-66.00 % Clay 17.18 8.51 0.72-41.28 Azimuth (degrees) 241 64 100-312 Slope (%) 25.4 3.19 6.00-37.00 r = 0.4996 - 0.739(DEPTH) - 0.281 (pH) + 0.222( AZIMUTH) (r2 = 0.432, P = 0.00) where: r = presence of piny on-juniper stands [1]. Soil depth was negatively correlated to favorable stand conditions. This may appear counter-intuitive, however other investigators found that when grass cover was heavy, seedlings probably could not compete for limited soil resources (Woodbury 1947; Aro 1971). These shallow, rocky sites provided conditions that result in little competition from herbaceous vegetation, leaving piny on-juniper able to survive. Soil pH was also negatively correlated, but that is not to say the more favorable sites were acid in nature; pH ranged from 6.7 to 8.8. Thus, a low pH for this study was one that approaches neutral. The data indicate that north to northwest faces were the most favorable exposures for pinyon- juniper stands which is consistent with similar results from the southern part of the state where pinyon exhibited higher cover values on northern exposures (Pieper & Lymbery 1987). The data support, that in the Capulin Volcano Field of northeastern New Mexico, sites with shallow, rocky soils, moderate pH and northern exposures provide suitable conditions for piny on-juniper to occur. 160 THE TEXAS JOURNAL OF SCIENCE-VOL. 69, NO. 2, 2008 Acknowledgments The authors wish to thank Andrew H. Feldman for providing critical review prior to publication. Literature Cited Aro, R. S. 1971. Evaluation of piny on- juniper conversion to grassland. J. Range Manage., 24:188-197. Bouyoucous, G. J. 1962. Hydrometer method improved for making particle size analysis of soils. Agron. J., 54:464-465. Bouyoucous, G. J. 1951. A recalibration of the hydrometer method for making mechanical analysis of soils. Agron. J., 43:434-438. Larson, F. R. 1980. Piny on-juniper 239. Pp. 116-117, in Forest cover types of the United States and Canada. F.H. Eyre ed. Soc. Am. For., 148 pp. McCurdy, G. 1994. Des Moines weather station 1916-1994. Western Regional Climate Center. Pieper, R. D. & G. A. Lymbery. 1987. Influence of topographic features on pinyon- juniper vegetation in south central New Mexico. Pp. 53-57, in proc. Proceedings Pinyon-Juniper Conference. Everett ed. USDA-Forest Service, Gen. Tech. Rep.INT-215 581 pp. SPSS. 2003. SPSS 12.0 user’s guide. SPSS Inc. Chicago, IL, 677 pp. Trauger, F. D. & T. E.. Kelly. 1987. Water resources of the Capulin topographic basin, Colfax and Union counties, New Mexico. Pp. 285-293, in Northeastern New Mexico. Lucas and Hunt eds. New Mexico Geolo. Soc., 354 pp. Woodbury, A. M. 1947. Distribution of pigmy conifers in Utah and northeastern Arizona. EcoL, 28:113-126. WDH at: David_Hacker@NMHU.edu THE TEXAS ACADEMY OF SCIENCE, 2008-2009 OFFICERS President President Elect: Vice-President: Immediate Past President: Executive Secretary: Corresponding Secretary: Managing Editor: Manuscript Editor: Treasurer: AAAS Council Representative: International Coordinator: Raymond C. Mathews, Jr., Texas Water Dev. Board William J. Quinn, St. Edward’s University Benjamin A. Pierce, Southwestern University Hudson R. DeYoe, University of Texas-Pan American Fred Stevens, Schreiner University Diane B. Hyatt, Texas Water Development Board Ned E. Strenth, Angelo State University Frederick B. Stangl, Jr., Midwestern State University John A. Ward, Brooke Army Medical Center James W. Westgate, Lamar University Armando J. Contreras, Universidad Autonoma de N.L. DIRECTORS 2006 Herbert D. Grover, Hardin-Simmons University Gary P. Garrett, Texas Parks and Wildlife Department 2007 Renard L. 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No part of this publication may be reproduced, stored in a retrieval system or transmitted, in any form or by any means, electronic, mechanical, recording or otherwise, without the prior permission of the Managing Editor of the Texas Journal of Science. THE TEXAS JOURNAL OF SCIENCE Volume 60, No. 3 August, 2008 CONTENTS Reproductive Cycle of the Common Rough-Scaled Lizard, Ichnotropis squamulosa (Squamata: Lacertidae) from Southern Africa. By Stephen R. Goldberg . . . . . . . . . 163 Secretory Vesicle Refilling and Toxin Production in Ollotis nebulifer (Coastal Plain Toad) and Anaxyrus speciosus (Texas Toad) following Transcutaneous Electrical Stimulation of the Parotoid Glands. By Randy L. Powell, M. Andres Soto, Romulo H. Gonzalez, Jr. and Jennifer R. Purviance . . . . . . . 171 Temperature Tolerance of Green Sunfish (Lepomis cyanellus). By Cheryl Cortemeglia and Thomas L. Beitinger . . 179 Evaluation of the Utility of the American College Testing Program Examination in Predicting the Probability of Success in Calculus I in Select Texas Universities. By Paul F. Messina . . . . . . . . . . . 189 Algal Community Structure of the East and West Flower Gardens, Northwestern Gulf of Mexico. By Erin M. Hill and Roy L. Lehman . . . . . 201 A Comparison of Mandibular Structure in Ruddy and Masked Ducks (Oxyura jamaicensis and Nomonyx dominicus). By Jack C Eitniear and Kent Ry lander . . . . . . . . . . 2 1 5 General Notes The Occurrence of Agardhiella ramosissima (Gigartinales) and Acanthophora spicifera (Ceramiales) in the Texas Coastal Bend. By Ryan L. Pikes and Roy L. Lehman . . . . . ......221 First Record of the Gray Fox {Urocyon cinereoargenteus) on Texas Barrier Islands. By Gerrad D. Jones and Jennifer K. Frey . . . . . 225 First Report of the Brown Alga Padina glabra (Ochrophyta: Dictyotales) from the Coast of Texas and the Gulf Of Mexico. By Michael J. Wynne.... . . . . . . . 227 Reproduction in the Siamese Leaf-Toed Gecko, Dixonius siamensis (Squamata: Gekkonidae) from Thailand. By Stephen R. Goldberg . . . . . . . 233 Recognition of Member Support . . . . . . ..239 Membership Application . . . . . 240 THE TEXAS JOURNAL OF SCIENCE EDITORIAL STAFF Managing Editor: Ned E. Strenth, Angelo State University Manuscript Editor: Frederick B. Stangl, Jr., Midwestern State University Associate Editor for Botany: Janis K. Bush, The University of Texas at San Antonio Associate Editor for Chemistry: John R. Villarreal, The University of Texas-Pan American Associate Editor for Computer Science: Nelson Passos, Midwestern State University Associate Editor for Environmental Science: Thomas LaPoint, University of North Texas Associate Editor for Geology: Ernest L. Lundelius, University of Texas at Austin Associate Editor for Mathematics and Statistics: E. Donice McCune, Stephen F. Austin State University Associate Editor for Physics: Charles W. Myles, Texas Tech University Manuscripts intended for publication in the Journal should be submitted in TRIPLICATE to: Dr. Frederick B. Stangl, Jr. TJS Manuscript Editor Department of Biology Midwestern State University Wichita Falls, Texas 76308 frederick.stangl@mwsu.edu Scholarly papers reporting original research results in any field of science, technology or science education will be considered for publication in The Texas Journal of Science. Instructions to authors are published one or more times each year in the Journal on a space-available basis, and also are available on the Academy's homepage at: www.texasacademyofscience.org AFFILIATED ORGANIZATIONS American Association for the Advancement of Science, Texas Council of Elementary Science Texas Section, American Association of Physics Teachers Texas Section, Mathematical Association of America Texas Section, National Association of Geology Teachers Texas Society of Mammalogists TEXAS J. OF SCI. 60(3): 1634 70 AUGUST, 2008 REPRODUCTIVE CYCLE OF THE COMMON ROUGH-SCALED LIZARD, ICHNOTROPIS SQUAMULOSA (SQUAMATA: LACERTIDAE) FROM SOUTHERN AFRICA Stephen R. Goldberg Department of Biology, Whittier College Whittier, California 90608 Abstract.-The reproductive cycle of the common rough-scaled lizard, Ichnotropis sqiiamulosa from southern Africa was studied from a histological examination of gonads. The smallest reproductively active male and female /. sqiiamulosa measured 47 mm and 58 mm SVL, respectively. Males began (spermiogenesis) sperm production and females began yolk deposition in February (summer). The reproductive cycle of 1. sqiiamulosa differs from that of other lacertid lizards from southern Africa which typically begin sperm production in late winter or early spring concurrent with the onset of yolk deposition. This difference in timing of reproduction may enhance survival of /. sqiiamulosa as its young appear in spring by which time neonates of other lacertid species are larger and have different dietary preferences. The common rough-scaled lizard, Ichnotropis squamulosa frequents arid and mesic savannah and ranges through Maputaland, Northern Cape, extreme south and central Mozambique, through Botswana, Zimbabwe and eastern Namibia to Angola and Tanzania (Branch 1998). It is a sit and wait predator that brumates during winter (Pianka 1971) and has a short life span of only eight to nine months (Broadley 1967; Schmidt 2001). There are previous reports on its reproductive biology (Fitzsimons 1943; Broadley 1967; 1974; 1979; Jacobsen 1987; Schmidt 2001). The purpose of this paper is to present additional information on the reproductive cycle of L squamulosa gathered from a histological examination of gonadal material from museum specimens. The reproductive cycle of /. squamulosa is compared with those of other lizards from southern Africa. Methods and Materials One hundred and one lizards were examined from the herpetology collection of the Natural History Museum of Los 164 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Angeles County, (LACM), Los Angeles, California. The sample consisted of 32 females (mean snout-vent length [SVL] = 56.3 mm ± 4.4 SD, range: 48-66 mm), 37 males (mean SVL = 53.0 mm ± 5.4 SD, range: 39-63 mm) and 32 juveniles (mean SVL = 33.4 mm ± 3.7 SD, range: 28-39 mm). Ichnotropis squamulosa were collected during 1969 and 1970 as part of an ecological study by Pianka (1971; 1986) or in 1972 and 1973. For histological examination, the left testis and epididymis were removed from males. The stages in the testicular cycle were identified. The left ovary was removed from females for histological examination to check for the presence of vitellogenesis (yolk deposition) and/or corpora lutea. Tissues were embedded in paraffin and cut into sections at 5 pm. Slides were stained with Harris hematoxylin followed by eosin counterstain (Presnell & Schreibman 1997). An unpaired Mest was used to compare /. squamulosa male versus female mean body sizes (SVL) and mean body sizes of November versus January juveniles (Instat vers. 3.0b, Graphpad Software, San Diego, CA). Material examined -^2imp\QS consisted of the following specimens of Ichnotropis squamulosa: BOTSWANA {n - 82) KGALAGADI DISTRICT, 11 km S. Tsabong (26°08’S, 22°28’E) 80272-80280, 80282, 80284, 80288, 80290-80292, 80295-80301, 80304, 80305, 80307-80318, 80320- 80322, 80326-80335, 80337-80342, 80345-80351, 80353, 80354, 80356-80358, 80360-80364, 80369, 80371-80374, 80376-80381. NAMIBIA {n= 12) OTOZONDJUPA REGION, 40 km WNW Grootfontein (19°34’S, 18°07’E) 77833, 77835, 77836; 30 km ENE Otavi (19°40’S, 17°24’E) 77879-77887. REPUBLIC OF SOUTH AFRICA {n = 7) NORTHERN CAPE PROVINCE, 31 km N, 100 km E. Upington (28°13’S, 22°16’E) 80265-80271. GOLDBERG 165 Results and Discussion Stages observed in the testicular cycle of /. squamulosa are given in Table 1: (1) Regression, seminiferous tubules contain mainly spermatogonia and primary spermatocytes; (2) Recrudes¬ cence, there is an increase in numbers of germ cells and cell divisions are noted. Secondary spermatocytes are abundant, a few spermatids may be present; (3) Late recrudescence, secondary spermatocytes and spermatids predominate; no sperm are present; (4) Early spermiogenesis, clusters of metamorphosing spermatids line portions of the lumina of the seminiferous tubules, occasional spermatozoa are seen; (5) Spermiogenesis, borders of seminiferous tubules are lined by rows of metamorphosing spermatids and spermatozoa are abundant. Monthly stages of the testicular cycle are in Table 1. While data is lacking from each month, a large February sample (w = 33) which contained 7/33 (21%) testes in recrudescence (germ cell renewal prior to the next period of sperm formation) and 21/33 (64%) in spermiogenesis indicates that L squamulosa probably commences sperm formation during that month. Spermiogenesis continues at least through April (Table 1). The smallest reproductively active male (spermiogenesis underway) measured 47 mm SVL (LACM 80337) and was from February. Females of L squamulosa were larger than males (unpaired /-test = 2.7, df= 61, P = 0.008). Three stages were observed in the ovarian cycle (Table 2): (1) No yolk deposition, ovary is quiescent; (2) Early yolk deposition, scattered vitellogenic granules are present; (3) Both corpora lutea from a previous clutch and concomitant yolk deposition for a subsequent clutch are present in the same female. It appears that the L squamulosa female population commences reproductive activity in February as two females (2/29, 7%) exhibited early yolk deposition (Table 2). However, Jacobsen (1987) reported 1 of 3 /. squamulosa females from January contained enlarged ovarian follicles. Yolk deposition continued through April (Table 2). Laying of eggs was underway by May as one female (LACM 80381, SVL = 66 mm) exhibited corpora lutea from a recent clutch. The same female was con- 166 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Table 1. Monthly distribution of reproductive conditions in the seasonal testicular cycle of 37 Ichnotropis sqiiamulosa from southern Africa. Values are the numbers of males exhibiting each of the five conditions. Month n Regression Recrudescence Late recrudescence Early spermio¬ genesis Spermio¬ genesis January 1 1 0 0 0 0 February 33 1 1 6 2 23 April 3 0 0 0 0 3 Table 2. Monthly stages in the ovarian cycle of 32 Ichnotropis squamulosa from southern Africa. Values are the numbers of females exhibiting each of the three conditions. Month n No yolk deposition Early yolk deposition Corpora lutea & yolk deposition February 29 27 2 0 April 2 1 1 0 May 1 0 0 1 currently undergoing yolk deposition for an additional clutch indicating 1. squamulosa females have the potential for producing two egg clutches in the same year. The smallest reproductively active 1. squamulosa (early yolk deposition) measured 58 mm SVL and was from February (LACM 80357). Juveniles were collected in November {n= 12, mean SVL = 32.3 mm ±4.1 S.D., range: 28-38 mm) and January {n = 20, mean SVL = 34.1 mm ±3.4 S.D.^ range: 28-39 mm). There was no significant size difference between the two months (unpaired ^test, t \ 3, df = 30, P = 0.21). Presumed neonates of 28 mm SVL were collected in both November and January. There is a report of hatchling 1. squamulosa measuring 28-30 mm SVL (Broadley 1979). Histological observations indicate that spermiogenesis likely commences in February concomitant with males acquiring breeding coloration (Schmidt 2001). The smallest male to undergo GOLDBERG 167 spermiogenesis in this study (47 mm SVL) was smaller than the minimum SVL (55 mm) in Schmidt (2001). Broadley (1979) reported a gravid female from 10 April. Eggs are laid in April-May with isolated cases as late as July (Jacobsen 1987). Pianka (1986) reported two 7. squamulosa females, each with a clutch of 4 eggs; a range of 10-12 eggs are deposited (Fitzsimmons 1943). This value of 58 mm SVL for the smallest breeding female in this study was identical with that of Schmidt (2001). Ichnotropis squamulosa has a short life span of only eight to nine months; young are bom October to November after a five month incubation period (Schmidt 2001). November collections reported herein consisted of only juveniles. Broadley (1974; 1979) similarly reported only juveniles in November and December. Hatchlings reach adult sizes in four to five months; breeding occurs in March and April with eggs being deposited in April and May (Broadley 1979; Schmidt 2001). Ichnotropis squamulosa is sympatric with Ichnotropis capensis however, the timing of their reproductive cycles differ so as to avoid competition for food (Broadley 1967, 1979; Jacobsen 1987). Ichnotropis capensis mate in October-December (Branch 1998) at which time 7 squamulosa have not yet reached maturity. The reproductive cycle of 7 squamulosa differs from that of other lacertid lizards from southern Africa which typically begin sperm production in late winter or early spring concurrent with the onset of yolk deposition (Goldberg 2006a; 2006b; 2006c; 2006g; 2006h; Goldberg & Robinson 1979; Nikosi et al. 2004). This difference in timing of reproduction may enhance survival of 7 squamulosa as its young appear in spring at which time neonates of other species of lacertids have reached sub-adult or adult size and have different dietary preferences. Aporosaura anchietae is an apparent exception as some reproduction occurs throughout the 168 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 year (Goldberg & Robinson 1979). This may, in part, be due to its ability to utilize seeds in their diet which are available year-round, (Robinson & Cunningham 1978). Conclusions Reproduction in 7. squamulosa is synchronous in that spermiogenesis and ovarian reproductive activity occur at the same time, however each take place in autumn whereas they occur primarily during spring in other lizards from southern Africa (Goldberg & Robinson 1979; Flemming 1994; Flemming & Bates 1995; Nkosi et al. 2004; Goldberg 2006a; 2006b; 2006c; 2006d; 2006e; 2006f; 2006g; 2006h). There are, however, exceptions in which male and female reproductive activity are asynchronous as in Cordylus giganteus (cf. Van Wyk 1991; 1995) and Pseudocordylus melanotus (cf Flemming 1993a; 1993b). In addition, females of the agamid lizard, Agama atra follow a condensed reproductive cycle in which ovarian activity occurs only in September to November (Van Wyk 1984). There is a report in a field guide (Branch 1998) that the sympatric congener, 7 capensis may produce two clutches in a year. However, additional work is needed to verify whether 7 squamulosa, in fact, may produce more than one egg clutch per year. Follicular atresia (spontaneous degeneration of oocytes with reabsorption of yolk) is common late in the reproductive season (Goldberg 1973). It is thus uncertain if the one 7 squamulosa female from the May sample with corpora lutea and concomitant yolk deposition would have produced a second egg clutch or undergone follicular atresia. Acknowledgements I thank C. Thacker (LACM) for permission to examine 7. squamulosa and J. Carlson and S. Kark (Whittier College) for assistance with histology. GOLDBERG 169 Literature Cited Branch, B. 1998. Field Guide to Snakes and other Reptiles of Southern Africa, 3*^^^ edn. Ralph Curtis Books Publishing, Sanibel Island, Florida, 399 pp. Broadley, D. G. 1967. The life cycles of two sympatric species of Ichnotropis (Sauria: Lacertidae). Zook Afr, 3:1-2. Broadley, D. G. 1974. Field studies on ‘annual lizards’ of the genus Ichnotropis. Rhodesia Sci. News, 8:309. Broadley, D. G. 1979. A field study of two sympatric ‘annual’ lizards (genus Ichnotropis) in Rhodesia. S. Afr. J. Zook, 14:133-138. Fitzsimons, V. F. 1943. The Lizards of South Africa. Transvaal Mus. Mem., Pretoria, South Africa, Mem. No. 1, 528 pp. Flemming, A. F. 1993a. The male reproductive cycle of the lizard Pseiidocordylus m. melanotus (Sauria: Cordylidae). J.Herpetok, 27:473-478. Flemming, A. F. 1993b. The female reproductive cycle of the lizard Pseiidocordylus m. melanotus (Sauria: Cordylidae). J. Herpetok, 27:103-107. Flemming, A. F. 1994. Male and female reproductive cycles of the viviparous lizard, Mabuya capensis (Sauria: Scincidae) from South Africa. J. Herpetok, 28:334-341. Flemming, A. F., & M. F. Bates. 1995. Male and female reproductive cycles of Bibron’s gecko Pachydactylus bibronii (Sauria: Gekkonidae) in the Free State province of South Africa. Afri. J. Zook, 109:409-422. Goldberg. S. R. 1973. Ovarian cycle of the western fence lizard, Sceloporus occidentalis. Herpetologica, 29:284-289. Goldberg, S.R. 2006a. Reproductive cycle of the Kalahari tree skink, Trachylepis spilogaster (Squamata: Scincidae) from southern Africa. Texas J. Sci., 58(4):291-298. Goldberg, S. R. 2006b. Reproductive cycle of the spotted sand lizard, Pedioplanis lineoocellata (Squamata: Lacertidae) from southern Africa. Texas J.Sci., 58(l):65-72. Goldberg, S. R. 2006c. Reproductive cycle of the Namaqua sand lizard, Pedioplanis namaquensis (Squamata: Lacertidae), from southern Africa. Afr. Zook, 41:147- 149. Goldberg, S. R. 2006d. Reproductive cycle of the Namib giant ground gecko, Chondrodactylus angiilifer (Squamata: Gekkonidae). Afr. Zook, 41:308-31 1. Goldberg, S. R. 2006e. Reproductive cycle of the striped legless skink, Typhlosauriis lineatiis (Squamata: Scincidae) from Southern Africa. Herpetok Bulk, 98:26-28. Goldberg, S. R. 2006f. Notes on the reproductive biology of the variegated skink, Trachylepis variegata (Squamata: Scincidae), from southern Africa. Bull. Chicago Herpetok Soc., 41:168-169. Goldberg, S. R. 2006g. Reproductive cycle of the spotted sand lizard, Meroles suborbitalis (Squamata: Lacertidae) from South Africa. Texas J. Sci., 58(3):255- 262. Goldberg, S. R. 2006h. Reproductive cycle of the bushveld lizard Heliobolus lugubris (Squamata: Lacertidae) from southern Africa. Salamandra, 42:151-154. 170 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Goldberg, S. R. & M. D. Robinson. 1979. Reproduction in two Namib Desert lacertid lizards (Aporosaura anchietae and Meroles cuneirostris). Herpetologica, 35:169-175. Jacobsen, N. H. G. 1987. Notes on reproduction in Ichnotropus squamulosa and interspecific competition with I. capensis (Reptilia, Lacertidae) in the Transvaal. J. Herpetol. Assoc. Afr., 33:13-17. Nikosi, W. T., N. J. L. Heideman, & J. H. Van Wyk. 2004. Reproduction and sexual size dimorphism in the lacertid lizard Pedioplanis burchelli (Sauria: Lacertidae) in South Africa. J. Herpetol., 38:473-480. Pianka, E. R. 1971. Lizard species density in the Kalahari Desert. Ecology, 52:1024-1029. Pianka, E. R. 1986. Ecology and Natural History of Desert Lizards. Analyses of the Ecological Niche and Community Structure. Princeton University Press. Princeton, New Jersey, 208 pp. Presnell, J. K., & M. P. Schreibman. 1997. Humason’s Animal Tissue Techniques, 5**^ Ed., The Johns Hopkins University Press, Baltimore, 572 pp. Robinson, M. D., & A. B. Cunningham. 1978. Comparative diet of two Namib Desert sand lizards (Lacertidae). Madoqua, 11:41-53. Schmidt, A. D. 2001. Autokologie und fortplanzungsbiologie von Ichnotropis squamulosa Peters, 1854 (Reptilia: Lacertidae). Salamandra, 37:157-174. Van Wyk, J. H. 1984. Ovarian morphological changes during the annual breeding cycle of the rock lizard Agama atra (Sauria: Agamidae). Navorsinge Nasionale Museum Bloemfontein, 4:237-275. Van Wyk, J. H. 1991. Biennial reproduction in the female viviparous lizard Cordyhis gigauteus. Amphib.-Reptil., 12:329-342. Van Wyk, J. H. 1995. The male reproductive cycle of the lizard, Cordylus gigauteus (Sauria: Cordylidae). J. Herpetol., 29:522-535. SRG at: sgoldberg@whittier.edu TEXAS J. OF SCI. 60(3):171-178 AUGUST, 2008 SECRETORY VESICLE REFILLING AND TOXIN PRODUCTION IN OLLOm NEBULIFER (COASTAL PLAIN TOAD) AND ANAXYRUS SPECIOSUS (TEXAS TOAD) FOLLOWWG TRANSCUTANEOUS ELECTRICAL STIMULATION OF THE PAROTOID GLANDS Randy L. Powell, M. Andres Soto, Romulo H. Gonzalez, Jr. and Jennifer R. Purviance* Department of Biological and Health Sciences, MSC 158 Texas A&M University, Kingsville, Texas 78363 and ^Department of Marine Biology, Texas A&M University 5007 Avenue U, Galveston, Texas 77553 Abstract.-Parotoid glands in two species of toads, Ollotis nebidifer and Anaxyrus speciosus were expressed using transcutaneous electrical stimulation. Toads were euthanized at various time stages following gland expression. Of the toads that had parotoid glands expressed, microscopic examination revealed the majority of the vesicles in the glands had emptied significantly with little to no contents remaining. General morphology of the vesicles was similar in both species with vesicles in A. speciosus somewhat smaller and more numerous than in O. nebulifer. Unexpressed vesicles were generally ovoid in shape with smooth cell walls. Emptied vesicles displayed numerous invaginations of the cell wall, resulting in the decrease of the vesicle volume. No gross injury or damage to the emptied vesicles or surrounding structures was noted. Hypertrophy of the inner (secretory) cell wall layer was evident at 24 h in both species. The majority of vesicles in both species had completely refilled and resumed normal size and shape within 5 to 7 days following gland expression. Amphibians possess two major types of integumentary glands that are widespread throughout the head, body, and limbs (Duellman & Tmeb 1985; Zug et al. 2001). The mucous glands, which are more abundant dorsally, secrete a clear slimy fluid that maintains a thin protective layer over the skin to prevent dehydration of the animal. The granular glands, which produce a biochemical mixture of poisonous or noxious secretions, tend to be more concentrated on the head and shoulder regions (Duellman & Trueb 1985; Zug et al. 2001). Granular glands are commonly aggregated into large macrogland structures (e.g., parotoid gland; see Tyler et al. 2001 for an in-depth analysis on the nomenclature and spelling) and are present in various degrees throughout the anurans and occur in some salamanders (Duellman & Trueb 1985). 172 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 3, 2008 Each granular gland consists of numerous alveoli or secretory vesicles that are surrounded by connective tissue (Hostetler & Cannon 1974; Cannon & Hostetler 1976; Alvarez et al. 2005). The vesicles produce and store toxic secretions and are the functional units of the granular glands. Microstructure of the vesicle is made up of a double cell layer consisting of an outer layer of myoepithelial cells that function as a contractile component and an inner layer of secretory cells that is continuous with the vesicle contents (Hostetler & Cannon 1974; Cannon & Hostetler 1976). Each vesicle has a duct that allows the biochemical mixture access to the skin surface (Hostetler & Cannon 1974; Cannon & Hostetler 1976; Toledo et al. 1992) where it acts as a defensive mechanism against predators (Duellman & Trueb 1985; Zug et al. 2001). Amphibian granular glands produce numerous biologically active compounds including; biogenic amines, peptides, proteins, bufadienolides, tetrodotoxins, and lipophilic alkaloids (see review Daly et al. 1987; Erspamer 1994). Because of the abundance of various biochemicals it contains and their application to human health, amphibian integument has generated a continual interest among researchers (Barthalmus 1994). The most common and “traditional” method of obtaining granular gland secretion has been a non-conservative technique that requires sacrificing the animal and subsequent mechanical and chemical processing of the skin (Roseghini et al. 1976; Erspamer et al. 1984; Erspamer et al. 1986; Roseghini et al. 1988; Roseghini et al. 1989; Daly et al. 2004). A much less common, but conservative method (the animal is not sacrificed) for the collection of amphibian granular gland secretion utilizes electrical stimulation (Conlon et al. 1999; Grant & Land 2002; Bevier et al. 2004). Electrical stimulation involves applying a low voltage of electric current (10-15 V) directly to the skin of the animal which causes contraction of the granular glands and ejection of their contents onto the skin surface where it can be collected (Tyler et al. 1992; Grant & Land 2002). Clearly, an advantage of using electrical stimulation is that the animal is not sacrificed and can be released back into its natural POWELL ET AL. 173 habitat, or if necessary, utilized again for serial extractions. However, currently there is no information available on the length of time required to refill or produce toxin in the granular glands after vesicle emptying. The present report describes a histological time study using electrical stimulation to express the granular gland contents in two species of anurans to examine vesicle emptying and refilling. Materlals and Methods Adult specimens (N=10 for each species) of the Coastal Plain Toad, Ollotis nebulifer and the Texas Toad, Anaxyrus speciosus (taxonomy follows Frost et al. 2006a, b) were utilized because of their abundance, large size, and prominent parotoid glands. All specimens were collected by hand at night within the city limits of Kingsville, Kleberg County, Texas USA and were housed individually in appropriately sized plastic Sterlite® containers which meet the requirements and guidelines for housing live amphibians (Pough 1991; National Academy Council 1997). Parotoid glands were expressed in live animals using a transcutaneous electrical stimulation device following Grant & Land (2002). Prior to electrical stimulation, the head and parotoid gland region was rinsed with deionized water; which aided in moistening the skin (to facilitate conduction of electrical current) and reduced contaminates in the collected glandular product. Each animal received three to four sessions of electrical massage directly to the parotoid glands for - 30 sec at 10 to 15 V. After each 30 second massage the expelled glandular contents was collected with a microspatula. The glandular secretion was frozen immediately and lyophilized for future analysis. Following gland expression, the animals were euthanized by standard approved methods (American Veterinary Medical Association 2001; Simmons 2002) at various time stages (control, 0 h, 12 h, 24 h, 2 d, 3 d, 5 d, 7 d) and the entire parotoid gland structure was formalized in situ with a 10% solution (3.7% formaldehyde) for one hour. Following formalin fixation the 174 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 3, 2008 parotoid glands were excised and soaked in Davidson’s solution (Kieman 2000) for 12 hours then rinsed with deionized water. The glands were placed in an automated tissue processor and dehydrated through a graded series of 70-100% ethanol and embedded in paraffin. Transverse and longitudinal sections were made for light microscopy examination at a thickness of 5 pm. The sections were mounted on glass slides and stained using standard hematoxylin and eosin methods (Sheehan & Hrapchak 1987; Kieman 2000). Results Size range for A. speciosus was 46-83 mm (total length), parotoid gland range; width 4.46-5.17 mm, length 7.34-12.17 mm. Size range for O. nebulifer was 74-102 mm (total length), parotoid gland range; width 5.48-6.79 mm, length 6.66-11.42 mm. The electrical stimulation procedure did not appear to cause any noticeable discomfort or injury to the animals. Parotoid glands from both species produced a cream colored viscous fluid that was easily collected. The product from A. speciosus was much thinner and noticeably more glutinous than in O. nebulifer. Microscopic examination of normal unexpressed vesicles in both A. speciosus and O. nebulifer were similar and appeared generally ovoid in shape with smooth cell walls (Fig. la & b). The unexpressed glands appeared full with morphology of the vesicles comparable in both species. In A. speciosus the vesicles were somewhat smaller than in O. nebulifer. Vesicle size in O. nebulifer was - 0.75 mm in diameter with vesicle size in A. speciosus - 0.5 mm in diameter. The contents of the full vesicles appeared homogenous in both species and cell morphology of the outer and inner layers of the vesicles were unremarkable. Following transcutaneous electrical stimulation, the majority of the storage vesicles in the glands had emptied significantly with little to no contents remaining. In addition, the general morphology of the vesicles had changed dramatically. In contrast to the ovoid shaped full vesicles, the emptied vesicles displayed numerous invaginations or infolding of the cell wall (Fig. Ic & d). The empty POWELL ET AL. 175 Fig. 1. Full unexpressed vesicles with smooth cell walls (a) Ollotis nebiiUfer (Coastal Plain Toad) (b) Anaxyrus speciosus (Texas Toad); Empty vesicles (EV) at 12 h showing numerous infolding of the cell wall, flattened angular shapes, and decrease volumes (c) Ollotis nebulifer (Coastal Plain Toad) (d) Anaxyrus speciosus (Texas Toad); Vesicles with marked hypertrophy of the inner (secretory) cell layer (SCL) with toxin production at 24 h (e) Ollotis nebulifer (Coastal Plain Toad) (f) Anaxyrus speciosus (Texas Toad). 176 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 3, 2008 vesicles displayed somewhat flattened angular shapes with marked decrease in volumes. There appeared to be no gross injury or damage to the cell walls, vesicles, or surrounding structures. Following vesicle emptying there was a marked hypertrophy of the inner (secretory) cell wall layer that was evident at 24 h in both species (Fig. le & f). In both A. speciosus and O. nebulifer^ there was a noticeable increase of fluid material within the vesicles after two to three days. Gradual refilling of the vesicles continued and normal “unexpressed” size and shape was resumed within five to seven days following gland expression. Discussion Transcutaneous electrical stimulation is a viable non-lethal method for the extraction and collection of anuran parotoid gland secretion. The process does not appear to cause any harmful effects to the animal as no gross or microscopic injury was evident during the procedure or revealed in histological examination. While there was a small difference in the average size of the vesicles between the two species examined, the time intervals for vesicle refilling was comparable. Empty vesicles refilled within seven days after the contents was extracted. However, it is important to note that the contents in the recently refilled vesicles (i.e., concentration of the toxins) were not analyzed and could possibly differ over time as the vesicles age or “mature”. The transcutaneous electrical stimulation method produced a relatively “clean” sample from the contents of the parotoid vesicles and does not require the time and expense of mechanical and chemical processing of the dried skin extraction method. Moreover, this conservative method would be preferred when dealing with rare or endangered species, it also allows for ontogenetic studies to be performed on the same animal, and would make feasible the use of anurans for venom production or farming similar to venom extraction facilities for venomous snakes. Acknowledgements Scientific Research Permit for collection of amphibians was issued by Texas Parks and Wildlife to RLP (SPR- 1205- 1546) and POWELL ET AL. 177 MAS (SPR-0102-188). All experiments with live animals were approved by Texas A&M University-Kingsville Institutional Animal Care and Use Committee (TAMUK # 2007-08-17). We would especially like to thank Frank Mullen and Michael Sean Walker, Department of Industrial Technology, Texas A&M University-Kingsville for their assistance in fabricating the transcutaneous electrical stimulation device. Literature Cited Alvarez, B. B., G. Delfino, D. Nosi & A. Terreni. 2005. Ultrastructure of poison glands of South American frogs: A comparison between Physalaemns albonotatus and Leptodactyhis chaquensis (Anura: Leptodactylidae). J. MorphoL, 263:247-258. American Veterinary Medical Association. 2001. 2000 Report of the AVMA panel on euthanasia. J. Am. Vet. Med. Assoc., 218:669-696. Barthalmus, G. T. 1994. Biological roles of amphibian skin secretions. Pp. 382-410 in Amphibian Biology 1: The Integument (H. Heatwole & G. T. Barthalmus, Eds.). Surrey Beatty and Sons, Australia, xii+418 pp. Bevier, C. R., A. Sormevend, J. Kolodziejek, N. Nowotny, P. F. Nielsen & J. M. Conlon. 2004. Purification and characterization of antimicrobial peptides from the skin secretions of the mink frog {Rana septentrional is). Comp. Biochem. Physiol. C., 139:31-38. Carmon, M. S. & J. R. Hostetler. 1976. The anatomy of the parotoid gland in Bufonidae with some histochemical findings. II. Bufo alvarius. J. Morphol., 148:137-159. Conlon, J. M., T. Halverson, J. Dulka, J. E. Platz & F. C. Knoop. 1999. Peptides with antimicrobial activity of the brevinin-1 family isolated from skin secretions of the southern leopard frog, Rana sphenocephala. J. Pept. Res., 54:522-527. Daly, J. W., C. W. Myers & N. \\diittaker. 1987. Further classification of skin alkaloids from Neotropical poison frogs (Dendrobatidae), with a general survey of toxic/noxious substances in the amphibia. Toxicon, 25:1023-1095. Daly, J. W., N. Noimai, B. Kongkathip, N. Kongkathip, J. M. Wilham, H. M. Garraffo, T. Kaneko, T. F. Spande, Y. Nimit, J. Nabhitabhata & T. Chan-Ard. 2004. Biologically active substances from amphibians: preliminary studies on anurans from twenty-one genera of Thailand. Toxicon, 44:805-815. Duellman, W. E. & L. Trueb. 1985. Biology of Amphibians. McGraw Hill, New York. 670 pp. Erspamer, V. 1994. Bioactive secretions of the amphibian integument. Pp. 178-350 in Amphibian Biology 1: The Integument (H. Heatwole & G. T. Barthalmus, Eds.). Surrey Beatty and Sons, Australia, xii+418 pp. Erspamer, V,, G. Falconieri Erspamer, G. Mazzanti & R. Ondean. 1984. Active peptides in the skins of one hundred amphibian species from Australia and Papua New Guinea. Comp. Biochem. Physiol. C., 77:99-108. Erspamer, V., G. Falconieri Erspamer, G. Mazzanti & R. Ondean. 1986. Active peptides in the skins of two hundred and thirty American amphibian species. Comp. Biochem. Physiol. C., 85:125-137. 178 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 3, 2008 Frost, D. R., T. Grant, J. Faivovich, R. Bain, A. Haas, C. Haddad, R. De Sa, A. Charming, M. Wilkinson, S. Donnellan, C. Raxworthy, J. Campbell, B. Blotto, P. Moler, R. C. Drewes, R. Nussbaum, J. Lynch, D. Green & W. Wheeler. 2006a. The amphibian tree of life. B. Am. Mus. Nat. Hist., 297:1-370. Frost, D. R., T. Grant & J. R. Mendelson III. 2006b. Ollotis Cope, 1875 is the oldest name for the genus currently referred to as Cranopsis Cope, 1875 (Anura: Hyloides: Bufonidae). Copeia, 2006(3):558. Grant, J. B. & B. Land. 2002. Transcutaneous amphibian stimulator (TAS): A device for the collection of amphibian skin secretions. Herpetol. Rev., 33:38-41. Hostetler, J. R. & M. S. Cannon. 1974. The anatomy of the parotoid gland in Bufonidae with some histochemical findings. 1. Bufo marinus. J. Morphol., 142:225-239. Kieman, J. A. 2000. Histological and Histochemical Methods: Theory and Practice, Third Edition. Harcort Publishers. 512 pp. National Academy Council. 1997. Guide for the Care and Use of Laboratory Animals. National Academy Press, Washington, DC. 140 pp. Pough, F. H. 1991. Recommendations for the care of amphibians and reptiles in academic institutions. Institute for Laboratory Animal Research. National Academy Press, Washington, DC. 33(4):S1-S21. Roseghini, M., V. Erspamer & R. Endean. 1976. Indole-, imidazole- and phenyl- alkylamines in the skin of one hundred amphibian species from Australia and Papua New Guinea. Comp. Biochem. Physiol. C., 54:31-43. Roseghini, M., G. Falconieri Erspamer & C. Severini. 1988. Biogenic amines and active peptides in the skin of fifty-two African amphibian species other than bufonids. Comp. Biochem. Physiol. C., 91:281-286. Roseghini, M., G. Falconieri Erspamer, C. Severini & M. Simmaco. 1989. Biogenic amines and active peptides in extracts of the skin of thirty-two European amphibian species. Comp. Biochem. Physiol. C., 94:455-460. Sheehan, D. C. & B. B. Hrapchak. 1987. Theory and Practice of Histotechnology, Second Edition. Battelle Press, Columbus, Ohio. 481 pp. Simmons, J. E. 2002. Herpetological Collecting and Collections Management, Revised Edition. Herpetological Circular No. 3 1 . Society for the Study of Amphibians and Reptiles, 153 pp. Toledo, R. C., C. Jared & A. Brunner Jr. 1992. Morphology of the large granular alveoli of the parotoid glands in toad {Bufo ictericus) before and after compression. Toxicon, 30:745-753. Tyler, M. J., T. Burton & A. M. Bauer. 2001. Parotoid or Parotid: on the nomenclature of an amphibian skin gland. Herpetol. Rev., 32:79-81. Tyler, M. J., D. J. M. Stone & J. H. Bowie. 1992. A novel method for the release and collection of dermal, glandular secretions from the skin of frogs. J. Pharmacol. Toxicol., 28:199-200. Zug, G. R., L. J. Vitt & J. P. Caldwell. 2001. Herpetology: An Introductory Biology of Amphibians and Reptiles, Second Edition. Academic Press. 630 pp. RLP at: randy.powell@tamuk.edu TEXAS J. OF SCI. 60(3): 179- 188 AUGUST, 2008 TEMPERATURE TOLERANCE OF GREEN SUNFISH {LEPOMIS CYANELLUS) Cheryl Cortemeglia and Thomas L. Beitinger Department of Biological Sciences University of North Texas, Post Office Box 5218 Denton, Texas 76201 Abstract -Mean (± SD) critical thermal maxima of green simfish (Lepomis cyanellus) acclimated from 10 to 35°C ranged from 33.5±0.83°C to 41.3±0.26°C while the critical thermal minima of individuals acclimated over the same temperatures extended from 0.2±0.28°C to 11.2±0.73°C. Temperature tolerance was significantly related to acclimation temperature and the latter accounted for a majority of variation in both lower (97%) and upper (98%) temperature tolerance. A measured upper lethal temperature of 4L3°C and estimated temperature tolerance polygon of 1270°C^ confirm that the green sunfish is among the most eurythermal native North American fish species. The tolerance polygon was partitioned into low and high temperature acclimation dependent areas of 384 and 205°C^, respectively, and when combined, essentially doubled the tolerance area from 618 to 1270°C“. The low temperature acclimation dependent area is approximately 1.4 times larger than the high temperature dependent area, suggesting that acclimation has a greater effect on tolerance of lower temperatures. The overall temperature tolerance ability of green sunfish may play an important role in this species ability to cope with global climate change. Green sunfish {Lepomis cyanellus) are native to the U.S. central plains between the Appalachian and Rocky Mountains, south to the Gulf Coast and north to the Great Lakes (Pflieger 1997). They are distributed throughout Texas and the U.S. with exceptions of Florida and parts of the northwest. The successful spread of green sunfish has been aided by their tolerance of physical/chemical variables (Smale & Rabeni 1995) which is an outcome of their origin and evolution within environmentally variable Great Plain streams. For example, water temperature in Great Plain streams reach 0°C in winter, and summer temperatures of 40° C have been reported (Matthews & Zimmerman 1990). Although a few studies (see Table 1) have presented upper temperature tolerances of green sunfish at selected acclimation temperatures; neither low temperature tolerance data nor a temperature tolerance polygon 180 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Table 1. CTMaxima (Mean + SD) of green sunfish. Entries include acclimation temperature, rate of temperature change during trials, test endpoint (Loss Of Equilibrium or Onset of Muscle Spasms) and source. Taccl (°C) AT (°C min' 9 Endpoint CTMax (°C) Source 25 0.3 LOE 37.4±0.53 Carveth et al. (2004) 30 0.3 LOE 40.2+0.50 Carveth et al. (2004) 26 0.017 LOE 37.9±0.75 Smale & Rabeni (1995) 10 1.0 LOE 31.1±1.45 Lutterschmidt & Hutchison (1997a) 10 1.0 OMS 34.2+2.06 Lutterschmidt & Hutchison (1997a) 20 0.3 LOE 35.8+0.69 Carrier & Beitinger (1988) 20 0.3 LOE 35.8+0.40 Carrier & Beitinger (1988) 20 0.3 LOE 35.9+0.56 Carrier & Beitinger (1988) across a range of acclimation temperatures has been published for this species. The objectives of this study were to quantify the acute upper and lower temperature tolerances of green sunfish acclimated to four constant temperatures. These data were used to generate an ecological temperature tolerance polygon following Bennett & Beitinger (1997) to more fully characterize the thermal tolerance of green sunfish. Materials and Methods Green sunfish were seined from a pond in Denton, Texas, transported to the laboratory in aerated containers and randomly allocated to one of four, well-aerated, glass aquaria containing dechlorinated tap water. Fish were held under a LD 12:12 photoperiod. Water temperature at the time of fish capture was approximately 25°C, and laboratory water temperature in each of the four holding aquaria (originally set to correspond with capture temperature) was changed 1°C d'^ until the desired acclimation temperature (10, 20 30 or 35°C) was attained. Haake® circulating thermoregulators (Models E-52 and D-1) were used to maintain 30°C and 35°C while the thermoregulators were used to heat CORTEMEGLIA & BEIJINGER 181 against constant cooling generated by a submerged cooler (Living Stream, Inc.) to maintain acclimation temperatures of 10 and 20°C. Once acclimated to 10, 20, 30 or 35°C, fish were held for 14 d before temperature tolerances were measured. The water quality of the holding tanks, measured twice weekly, was within ranges that should have produced no adverse effects to the fish (conductivity 312-700 pmhos cm"\ dissolved oxygen 6,4- 11.0 mg L I, pH 7.4-8. 0, ammonia (O.O-O.O mg L"^), and nitrite (0- 0.0 mg L‘^). Fish were fed Tetramin® tropical flake food twice daily during acclimation, but food was withheld one day prior to temperature trials. Daily water exchanges (5-25%) by siphon removed excess food and feces without changing water temperature. Critical thermal methodology, CTM, (Cox 1974) was used to estimate lower (CTMinima) and upper (CTMaxima) temperature tolerance of green sunfish acclimated to 10, 20, 30, and 35°C. The most recent reviews of this technique are provided by Lutterschmidt & Hutchison (1997b) and Beitinger et al. (2000). Specifics of the experimental CTM used in this research are identical to those of Cortemeglia & Beitinger (2005). During trials, the rate of temperature change was 0.3°C mm \ and water temperature was recorded each minute within + 0.1 °C with an ASTM calibrated mercury thermometer. Final loss of equilibrium was the endpoint criterion during CTMaxima trials and lack of response to prodding with a glass rod was the endpoint for CTMinima trials. Different test endpoints were required since fish exposed to increasing or decreasing temperature respond differently. Once each fish reached the endpoint, the temperature was recorded, the fish was removed from the CTM chamber, returned to its acclimation temperature, and assessed for survival for 24 hours. Each fish experienced either a heating or cooling trial, i.e., each fish was tested only once. Subsequently, each fish was weighed (wet, ± 0.05 g) and measured (total length, ±0.1 cm). Trials were performed between 1400 and 182 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 1800h to minimize possible diel fluctuations in temperature tolerance (Hutchison 1976). The CTMaximum and CTMinimum for each acclimation group were calculated as the arithmetic mean temperature at which the endpoint was observed. Thermal scope was calculated at each acclimation temperature and is defined as the arithmetic difference between CTMaximum and CTMinimum at a specific acclimation temperature. A dynamic temperature tolerance polygon combining both measured and extrapolated tolerance was developed following Bennett & Beitinger (1997). The polygon was partitioned following Beitinger & Bennett (2000) into three distinct zones: an upper acclimation temperature dependent zone that represents heat gain via acclimation, a lower temperature dependent zone that represents gain in low temperature tolerance and a middle tolerance zone that is independent of acclimation temperature. One-Way ANOVA followed by Student - Newman -Keuls (SNK) multiple comparison analyses tested for significant differences in mean CTMaxima and CTMinima at the four acclimation temperatures, and regression analyses determined if a linear relationship existed between acclimation temperature and either lower or upper temperature tolerance. Finally, simple linear regression analyses were used to determine if lower or upper temperature tolerances were related to either fish length or weight in each of the eight trials. All statistical decisions were based on an a of 0.05. Results Increasing acclimation temperature resulted in green sunfish gaining heat tolerance and losing “cold” tolerance. The Critical Thermal Maxima, CTMaxima, (mean ± SD) of green sunfish acclimated from 10°C to 35°C ranged from 33.5+0.83 to 41.3±0.26°C (Table 2). Mean CTMaxima were significantly different (One-way ANOVA, p < 0.0001) at all four acclimation temperatures (SNK multiple range analysis). Also a highly significant linear relationship (r^ = 0.96, p < 0.0001) exists: CTMaxima (°C) = 30.1 + 0.32(Taccl, °C). CORTEMEGLIA & BEITINGER 183 Table 2. Mean ( ± SD) critical thermal maxima and critical thermal minima and thermal scope of green sunfish at four acclimation temperatures. Sample size was ten in each group except the 35°C-CTMaximum group, which was 9. The thermal scope equals the difference between CTMaximum and CTMinimum at a particular acclimation temperature. Taccl(°C) CTMaxima (°C) CTMinima (°C) Thermal Scope (°C) 10°C 33.5+0.83 0.2+0.28 33.3 20°C 36.2±0.59 3.2+0.84 33.0 30°C 39.9±0.67 8.6+0.24 31.3 35°C 41.3±0.26 11.2+0.73 30.1 Mean (± SD) CTMinima of green sunfish acclimated to the same temperatures extended from 0.2± 0.28 to 11. 2± 0.73°C. One-way ANOVA (p < 0.0001) and SNK multiple range analyses determined that mean CTMinima also were significantly different at all four acclimation temperatures. A highly significantly (r = 0.97, p< 0.0001) relationship exists: CTMinima (°C) = -4.8 + 0.45(Taccl,°C). Thermal scopes ranged from 30.1 to 33.3°C. The area of the green sunfish ’s dynamic (ecological) temperature tolerance polygon encompassed 1270°C^ (Figure 1). The upper and lower acclimation dependent tolerance areas were 268®C and 384°C , respectively. Combined (652°C^) they are slightly larger than the area of the acclimation independent zone, 618°C^. The lower acclimation dependent zone was 1 .43 times larger than the upper acclimation dependent zone. Fish were between 3.9 and 9.0 cm long and weighed between 0.5 and 11.2 g, and no linear relationship was found between either length or weight and temperature tolerance for any of the eight trials, i.e., fish size did not significantly influence temperature tolerance in this study. 184 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 O o O 0 10 20 30 40 50 Acclimation Temperature (^C) Figure I. Ecological temperature tolerance polygon for green sunfish. The area (6I8T^) of the middle zone, B, represents temperature tolerance that is independent of acclimation temperature. Zones A and C represent upper (205°C^) and lower (384°C^) temperature tolerance areas produced by acclimation. Combined these two acclimation dependent areas account for slightly more than one-half of the entire ecological temperature polygon of 1270 Discussion Owing to its multiple effects (Fry 1947) on poikilotherms such as fish, temperature has been termed the abiotic master factor (Brett 1971). The most dramatic abiotic effect is lethality, i.e., too little or too much heat can kill an organism. Temperature tolerance determinations quantify the thermal limits of a species, offer insights into a species’ physiology, biology and ecology, and provide data to model possible effects of global climate change. Data generated by this research suggest that the success of green sunfish within Great Plains streams and their widespread dispersion throughout North America has been aided by their temperature tolerance ability. The large thermal scopes, extensive temperature tolerance polygon and highest CTMaximum (4L3°C) confirm that CORTEMEGLIA & BEITINGER 185 the green sunfish is both eurythermous and heat tolerant. A comprehensive review of temperature tolerances of North American freshwater fishes, indicated that only 22 species have a reported CTMaximum equal to or greater than 40°C. Of these only two are centrarchids: bluegill, Lepomls macrochirus, 41.4°C (Holland et al. 1974 and Murphy et al. 1976) and largemouth bass, Micropterus salmoides, 40.9°C (Fields et al. 1987). The mean CTMinimum of green sunfish acclimated to 10°C was 0.2°C. Loss of equilibrium was not observed in 7 of the 10 fish during this trial and the CTMinima of those seven individuals was recorded as 0.05°C (the lowest temperature attainable with the available cooling system). If acclimated to temperatures below 10°C, it is hypothesized that their low temperature tolerance allows green sunfish to survive the coldest temperatures attainable in freshwaters. The largest thermal scope and dynamic thermal tolerance zone reported for a North American fish are 34°C and 1470 for the sheepshead minnow, Cyprinodon variegatus, by Bennett & Beitinger 1997. Comparable variables for green sunfish are 33.3°C and 1270°C^. The 14% difference in thermal tolerance polygon area is explained by the 45.1°C CTMaximum for sheepshead minnow. A temperature tolerance polygon not only pictorially describes the temperature tolerance ability of a species, Beitinger & Bennett (2000) developed a geometrical approach to quantify the effect of acclimation on temperature tolerance. In it, a species temperature tolerance polygon is partitioned into three distinct zones: an upper acclimation temperature dependent zone which represents head gain via acclimation, a lower acclimation temperature dependent zone which represents gain of “cold” tolerance, a middle tolerance zone that is independent of acclimation temperature, termed the intrinsic temperature tolerance zone. This technique was applied first to polygons generated from incipient lethal temperature (ILT) 186 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 methodology for 21 species of fish (Beitinger & Bennett 2000). Although this is the first application of this partitioning technique to CTM data, our results for green sunfish are consistent with those for ILT generated tolerance data. First, the combined areas of the acclimation dependent zones comprised approximately one-half (51.2%) of the total tolerance polygon. This compares favorably with the mean 51.1% for the 21 species in the original study. Second, the lower acclimation dependent zone was larger (1.4 times) than the upper acclimation dependent zone. This suggests that acclimation plays a larger role in tolerance of low rather than high temperatures. Finally, applying a regression model to predict polygon area solely as a function of ultimate upper temperature yielded a predicted polygon area for green sunfish of 1258.4®C^ This is within 99% of this study’s empirical determined value of 1270°C . In toto, these finding suggest that this approach can be applied to CTM as well as ILT generated data, and reaffirms claims (see reviews by Lutterschmidt & Hutchison 1997b and Beitinger & Bennett 2000) that acclimation plays a major role in a species’ temperature tolerance. In addition to enhancing the area of the temperature tolerance polygon, variation in acclimation temperature accounted for 96% and 97% of the variation in upper and lower temperature tolerance, respectively. For each 1°C increase in acclimation temperature, green sunfish gained 0.32°C in heat tolerance, and for each decrease of 1°C in acclimation temperature, the mean CTMinimum decreased by 0.45°C. Species such as the green sunfish with large slopes relating acclimation temperature and temperature tolerance should be able to adjust lethal endpoints quickly when confronted with exposures to extreme temperatures (Beitinger & Bennett 2000). In conclusion, depending on its previous acclimation temperature, green sunfish are able to withstand temperatures from 0 to 41.3°C. This high degree of eurythermicity appears to have arisen from evolving within thermally labile streams of the U.S. CORTEMEGLIA & BEITINGER 187 Great Plains, and may allow this species to survive extreme temperatures expected by ongoing global climate change. Acknowledgements The authors greatly appreciate the technical assistance of Charles Pittman, Steve Timmons and Greta Bolin, and the help of Cynthia Thomas in collecting green sunfish. Research was conducted under the University of North Texas Institutional Animal Care and Use Committee (0410). Literature: Cited Beitinger, T. L. & W. A. Bennett. 2000. Quantification of the role of acclimation temperature in temperature tolerance of fishes. Env. Biol. Fish., 58:277-288. Beitinger, T. L., W. A. Bennett & R. W. McCauley. 2000. Temperature tolerances of North American freshwater fishes exposed to dynamic changes in temperature. Env. Biol. Fish., 58:237-275. Bennett, W. A. & T. L. Beitinger. 1997. Temperature tolerance of sheepshead minnow, Cyprinodon variegatus. Copeia, 1997:77-87. Brett, R. 1971. Energetic responses of salmon to temerature. A study of some thermal relations in the physiology and freshwater ecology of sockeye salmon {Oncorhynchus nerka). Amer. Zook, 1 1:99-1 13. Carrier, R. & T. L. Beitinger. 1988. Resistance of temperature tolerance ability of green sunfish to cadmium exposure. Bull. Environ. Contam. Toxicol., 40:475- 480. Carveth, C. J., A. Widmer, S. A. Bonar & W. Matter. 2004. Estimation of acute upper lethal water temperature tolerances of native Arizona fishes. A report submitted to the Water Resources Research Center, University of Arizona, 19 pp. Cortemeglia, C. & T. L. Beitinger. 2005. Temperature tolerances of wildtype and red transgenic zebra danios Danio rerio. Trans. Am. Fish. Soc., 134:143 1-1437. Cox, D. K. 1974. The effects of three heating rates on the crtical thermal maximum of bluegill. Pp. 158-163, in: J.W. Gibbons & R.R. Sharitz (ed.) Thermal Biology. Nat. Tech. Informa. Serv., Springfield, 686 pp. Currie, R. J., W. A. Bennett & T. L. Beitinger. 1998. Critical thermal minima and maxima of three freshwater game-fish species acclimated to constant temperatures. Env. Biol. Fish., 51:187-200. Fields, R., S. S. Lowe, C. Kaminski, G. S. Whitt & D. P. Philipp. 1987. Critical and chronic thermal maxima of northern and Florida largemouth bass and their reciprocal Fi and F2 hybrids. Trans. Amer. Fish. Soc., 1 16:856-863. Fry, F. E. J. 1947. Effects of the environment on animal activity. Univ. Toronto Studies in Biol., Series No. 55. Publ. Ont. Fish. Res. Lab., 68:1-62. 188 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Holland, W. E., M. H. Smith, J. W. Gibbons & D. H. Brown. 1974. Thermal tolerances of fish from a reservoir receiving heated effluent from a nuclear reactor. Physiol. Zool., 47:110-118. Hutchison, V. H. 1976. Factors influencing thermal tolerance of individual organisms. Pp. 10-26 in G. W. Esch and R. W. McFarlane, editors. Thermal Ecology IT ERDA (Energy Research and Development Administration) Symposium Series, CONE 752425, 404 pp. Lutterschmidt, W. E & V. H. Hutchison. 1997a. The critical thermal maximum: data to support the onset of spasms as the definitive end point. Can. J. Zool., 75:1553- 1560. Lutterschmidt, W. I. & V. H. Hutchison. 1997b. The critical thermal maximum: history and critique. Can. J. Zool., 75:1561-1574. Matthews, W. J. & E. G. Zimmerman. 1990. Potential effects of global warming on native fishes of the southern Great Plains and the Southwest. Fisheries 15:26-32. Murphy, J. C., C. T. Garten, Jr., M. H. Smith & E. A. Standora. 1976. Thermal tolerance and respiratory movement of bluegill from two populations tested at different levels of acclimation temperature and water hardness. Pp. 145-147, in: G. W. Esch & R. W. McFarlane (ed.) Thermal Biology II. Nat. Tech. Inform. Serv., Springfield, 404 pp. Pflieger, W. L. 1997. The Fishes of Missouri, Revised Edition. Missouri Department of Conservation, Jefferson City, Missouri, 372 pp. Smale, M. A. & C. F. Rabeni. 1995. Hypoxia and hyperthermia tolerances of headwater stream fishes. Trans. Am. Fish. Soc., 124:698-710. TLB at: beitingr@unt.edu TEXAS J. OF SCI. 60(3): 189-200 AUGUST, 2008 EVALUATION OF THE UTILITY OF THE AMERICAN COLLEGE TESTING PROGRAM EXAMINATION IN PREDICTING THE PROBABILITY OF SUCCESS IN CALCULUS I IN SELECT TEXAS UNIVERSITIES Paul F. Messina Department of Mathematics University of the Incarnate Word San Antonio, Texas 78209 Abstract -The efficacy of the American College Testing (ACT) Program examination in predicting success in a standard Calculus I course is analyzed. Using an ex-post facto design and the method of binary logistic regression, a significant relationship exists ACT scores and Caleulus I final grades of the 981 students examined during this study. Specifically, the data indicate that the ACT mathematics score was the most important predictor of success in Calculus I. Other covariates included in the final analysis were the ACT English score and student gender. Although ethnicity was considered in the analysis, it was found to be statistically insignificant to the final results. Colleges and universities participating in this study included Abilene Christian University, Angelo State University, Lamar University, Texas State University, and the University of Texas at San Antonio. Program planning and curriculum development require an accurate assessment of the student entering the educational system. A fundamental concern is the development of a program that is sensitive to accurate placement and evaluation. Students are traditionally asked by institutions to submit their American College Testing (ACT) Program scores, or other similar scores, to help institutions determine admission status. Scores are also often used to determine the initial placement of students in certain courses. This study is concerned with the relationship between the ACT Mathematics score and the grade earned in Calculus I. The primary research question was, for students enrolled at the institutions considered in this study for a five-year period, who had completed Calculus I and taken the ACT, is there a relationship between the final grade in Calculus I and the score on the ACT Mathematics test? Secondary analysis was conducted on several 190 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Other factors including the ACT composite score, ACT science reasoning score, ACT reading score, ACT English score, student ethnic group, student gender group, and academic institution. Justification for using standardized tests as a device for placement and predicting achievement of new college students generally has centered on the possible effects of poor placement with respect to their motivation and performance (Dunn 1966). The ACT program scores are used throughout the United States as well as in several other countries as an indicator of potential academic success at the tertiary level. It differs from its closest competitor, the SAT in a number of ways. The SAT is more popular in the eastern and far western United States, whereas the ACT is used more frequently in the Midwestern, southern, and southwestern United States. In addition to the verbal and mathematical sections found on the SAT, the ACT emphasizes reading and science reasoning. A standard scale is used for reporting total test scores on the ACT ranging from 1 (lowest) to 36 (highest). The overall mean standard score on the ACT Mathematics Test for entering college freshman during the course of the study was 20.7. Internal consistency reliability data of the ACT Mathematics Usage Test for students considered in this study was 0.91. The overall mean standard score on the ACT English, Math, Reading and Science Reasoning Test for entering college freshman during the course of this study was 20.4, 21.7, 21.3 and 21.0 respectively with an overall composite score of 21.0. According to these results, the ACT Assessment Test has remained stable over the course of this study. The selection of Calculus I as the specific course was predicated on the belief that undergraduate mathematics courses (at the level of Calculus and above) have been looked upon as an insurmountable barrier by many students. Students who would like to pursue study in the areas of medicine, dentistry, optometry, engineering, and computer science as well as other disciplines may fail to enter that career track because of a poor score in Calculus I (Lowe, 1981). Since mathematics is the primary tool for problem MESSINA 191 solving in many academic areas, a strong relationship between a student’s ACT Mathematics score and the final grade earned in Calculus I may provide evidence to support a change of the Mathematics curriculum at the secondary school level. In this study, the ACT Mathematics Usage Test was selected as the measuring device to predict success in Calculus I at five selected institutions of higher learning within the State of Texas. While there have been extensive studies of a related nature, few have looked at a student’s ACT score and their final grade in Calculus I, and none have used the method of binary logistic regression to evaluate the relationship. Binary logistic regression is appropriate in this instance because the response variable takes on only two values: success or failure. Ordinary least squares regres¬ sion assumes the response variable is continuous, so would be inappropriate in this instance. For an overview of logistic regres¬ sion, see Peng & So (2002). Hudson & Mclntire (1977) reported a weak correlation between performance in introductory non-Calculus physics and mathematics skills at the beginning of the course, and concluded that such skills alone are not sufficient to guarantee success in physics. In a more comprehensive study, Hudson & Rottman (1981) found a signifi¬ cant correlation {r-square = .175) between scores on a pre-course test of algebraic and trigonometric skills and performance in non- Calculus, pre-professional physics for those who completed the course. Wollman & Lawrenz (1984) subsequently verified the correlation between mathematics skills at the time of entry into the course and achievement in non-Calculus physics for those who completed the course. Another previous study, which was con¬ ducted at Pima College, showed that there is a weak correlation (r- square = .106) between the Nelson-Denny (N-D) Reading Test and the grade which was earned in introductory non-calculus-based physics (ladevaia 1989). 192 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Calculus I, although fundamental in nature, draws heavily upon a student’s mathematical preparation at the secondary school level. Thus, working from the assumption that the ACT Mathematics Assessment test is the best indicator of the mathematical preparation of the student, there should exist a moderately strong correlation between the ACT Mathematics Assessment score and the grade earned in Calculus I at five institutions of higher learning within the State of Texas. Another, conducted at Southern Illinois University at Edwardsville, focused on the differential effects of initial Chemistry course placement as a function of ACT Mathematics scores and secondary school rank-in-class percentile (Reiner 1971). It was found that students who were initially placed in the first course of the sequence would be expected to attain higher grade point averages if their rank-in-class was in the middle or lower, regardless of the student’s ACT Mathematics score. Data Collection and Statistical Analysis Data used in the analysis were obtained from each university’s Office of Institutional Research, and included results from 981 students attending five different universities in the state of Texas (Table 1) including Abilene Christian University (ACU), Angelo State University (ASU), Lamar University (LU), Texas State University (TSU) and the University of Texas at San Antonio (UTSA). Each university represented a mix of public, private, large, small, suburban, and rural universities as well as both competitive admissions and open admissions universities. Each of the schools was randomly selected from colleges and universities in Texas and each student who took Calculus I during the study timeline in each of these schools were included. Several variables were included in the analysis: ACT composite score (COMP), ACT English score (ENGL), and mathematics score (MATH) all with a continuous value range between 1 and 36; student gender (GEND) which was a dichotomous variable where 0 indicated male and 1 indicated female; and Calculus I final grade MESSINA 193 Table 1. Total university descriptive statistical data for five Texas universities. Classification N Abilene Christian University Angelo State University Lamar University Texas State University University of Texas at San Antonio African American/ Black 38 7 5 6 16 0 Asian/Pacific Islander 22 6 6 0 7 3 Hispanic 158 5 46 1 91 15 Caucasian 757 296 254 23 173 11 Other 6 0 0 3 3 0 Male 609 103 200 11 277 18 Female 372 63 111 22 165 11 Table 2. Average ACT scores of students in the population tested from 1998-2002. Students English Math Reading Science Composite 981 22.5 24.3 23.4 23.5 23.4 Table 3. Variables in the logistic regression model. B S.E. Wald Df Sig. Exp(B) ACTENG .057 .023 6.398 1 .011 1.059 ACTMAT .106 .028 14.558 1 .000 1.112 GENDER -.351 .145 5.888 1 .015 .704 Constant -.755 1.696 .198 1 .656 .470 (GRAD) which was also dichotomous for this study where 1 indicated successful completion of Calculus I (a final grade of A, B or C) and 0 indicated unsuccessful completion of Calculus I (a final grade of D, F or W). Student ACT data is shown below (Table 2). The models that were successfully fitted will be discussed. All data was analyzed using SPSS version 14.0. The initial analysis considered all variables for which data were available. As a result, it was easy to establish a baseline from which variables shown to be non-significant could be eliminated. The roles of the parameters in the model may be summarized by observing the variables in the logistic regression model (Table 3). In the table, B is the estimated coefficient. If the parameter is significant (jo<0.05) then the parameter is useful to the model. 194 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Table 4. Calculus I successful completion classification table. Non-successful Completion Successful Completion Percentage Correct Predicted Outcome Non-successful Completion 177 227 43.8 Actual Outcome Successful Completion 100 642 86.5 Totals 277 876 1= 1153 The next step involved determining how success was defined. An unfavorable response was coded as a 0, while a favorable response was coded a 1 . In this study, a final grade of A, B, or C was classified as successful while a grade of D, F, or W was classified as unsuccessful. A positive coefficient indicated that increasing values of the variable resulted in an increased likelihood of a favorable response. Initially the overall significance of the model was determined. The P'Value was < 0.001 for the step, block, and model, which indicates a model that is statistically significant. The Cox & Snell R-Square statistic had a value of .146 and the Nagelkerke R-Square statistic was .201. This indicates that the model performed in an acceptable manner. The classification table (Table 4) shows that successful completion of Calculus I could be predicted 86.5% of the time. The overall prediction ability average was 71.5%. In order to visually confirm the results, the data set was transformed into graphical output. A cross-tabulation of final grade in Calculus I and ACT Math score was performed. The data in the resulting frequency table was used to calculate the pass rate. A scatter plot was then created with the ACT Math score on the x-axis and the pass rate on the y-axis (Figure 1). MESSINA 195 Figure L Calculus I pass rate vs. ACT mathematics score. 1.1 1.0 .9 .8 ra K S .6 I -5 3 _o ro O -4 10 20 30 40 ACT Mathematics Score Figure 2. Calculus I moving average pass rate vs, ACT mathematics score. ACT Mathematics Score The graph shows a positive correlation of the data. After smoothing the data a scatter plot was created with the moving average pass rate on the y-axis and the ACT Math score on the x- axis (Figure 2). 196 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Figure 3. Final overlay scatter plot for ACT mathematics score. ACT Mathematics Score Figure 4. Calculus I pass rate vs. ACT English score. 1.2 1.0 CD .4 0 10 20 30 40 ACT English Score A binary logistic regression was performed with grade as the dependent variable and ACT Math score as the covariate. An overlay scatter plot was created which specified ACT Math score and pass rate as a pair and ACT Math score and the predicted probabilities as the second pair with the ACT Math score defined on the x-axis each time (Figure 3). This graph shows a significant relationship between the ACT Math score and the final grade earned in Calculus L The ACT Math score is a crucial factor in this study and shows a relationship that makes sense. MESSINA 197 Figure 5. Calculus I moving average pass rate vs. ACT English score. 1.0 -| - (U 10 20 30 40 ACT English Score Figure 6. Final overlay scatter plot for ACT English score. ACT English Score The same statistical analysis technique was used for the ACT English scores yielding the following results (Figures 4-6). Based upon the statistical analysis a binary logistic regression model was fit that includes the dependent variable GRAD and the independent variables MATH, ENGL, and GEND. The general logistic regression model and corresponding predictive equation took the following form: 198 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 In = « + A Pi {math) + (engJ) Therefore, ^ci+P\ ( getid)+P2 ( niath)+p^[engl) \ + ^^P[gendYPPmathYp^{engl) where a is the intercept or constant, and and/?3 represent the partial regression coefficient for jc,,x2, andx3 respectively, holding the remaining predictors constant. A sample calculation is included below for reference. Consider a male university student with an ACT Mathematics score of 23 and an ACT English score of 21. This student’s probability of success (e.g., obtaining a grade of C or above) is obtained as follows. ^a+P^(gend)+P2 {math)+ P^engl) ^ 1 + (gend)+P20>Jath)+P2{engl) ^-0.755+(-0.351)(0)+(0.106)(23)+(0.057)(21) 2 ^-0.755+(-0.351)(0)+(0.106)(23)+(0.057)(21) 2 88 ;r = -^^ = 0.9468 Therefore, the student’s probability of success is 0.9468. If we consider a female student with the same ACT characteristics, the following is obtained. ^a+PPgend)+P2imath)+P2^engl) I P^igend)^- P2{math)^ P^iengl) ^-0.755+(-0.351)(l)+(0.106)(23)+(0.057)(21) Y ^-0.755+(-0.351)(l)+(0.106)(23)+(0.057)(21) ;r = ^2.529 = 0.9261 MESSINA 199 In this case, the female student’s probability of success is 0.9261. While this equation was quite successful in this study it must be noted that the results presented can only “safely” extrapolate to the universities sampled, however, similar studies at other schools would be easy to do and are encouraged. Results and Discussion Binary Logistic Regression was conducted to determine which independent variables (MATH, ENGL, COMP, GEND, ACU, ASU, LU, TSU, and UTSA) were predictors of the final grade earned in Calculus I (GRAD). The statistics for overall model fit indicate a fairly good fitting model. The overall model correctly classified 86.5% of the cases. The analysis suggests that there is a positive relationship between the covariates under consideration and a student’s final grade in Calculus I within the State of Texas. As predicted earlier, it was found that the strongest relationship occurred between a student’s ACT mathematics score and the final grade earned in Calculus 1. In fact, compared to previous studies that have been done, the results obtained here are as good as or better than the previous research results. This study has several shown that a student’s predicted success in a standard Calculus I course in the State of Texas is directly related to their success on the American College Testing Program Examination (ACT). Colleges and universities can use these results to correctly place incoming students into an appropriate sequence that will enhance student’s chances for success. This will provide the student with best possible chance of being successful in collegiate level mathematics. Literature Cited ACU’s History. Abilene, Texas: Abilene Christian University. Retrieved May 30, 2002 from the World Wide Web: http://www.acu.edu/aboutacu/history.html Adams, L. LaMar, H. Bruce Higley, Leland H. Campbell. 1976. Statistical comparison of entrance prediction equations using ACT or SAT scores, or both. (ERIC Document Reproduction Service No. EJ 131 238). 200 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 History of ASU. Angelo, Texas: Angelo State University. Retrieved June 1, 2002 from the World Wide Web: http://www.angelo.edii/visitors/history_of_asu.htm Callahan, S. 1993. Mathematics placement at Cottey College. (ERIC Document Reproduction Service No. ED 373 813). Dunn, J. E. 1966. A study of the University of Arkansas mathematics entrance exam as a placement device. Journal of Experimental Education, 34, 62-68. Gagne, R. 1988. Principles of instruction design. Holt, Rinehart and Winston, Inc. (3""^ Edition), 321 pp. Hudson, H. T. & W. R. McIntyre. 1977. Correlation between mathematical skills and success in physics. American Journal of Physics, 45, 470-471. Hudson, H. T. & R. M. Rottman. 1981. Correlation between performance in physics and prior mathematics knowledge. Journal of research in science teaching, 18, 291-294. Jenkins, J. H. 1990. Placement practices for calculus in two-year colleges. (ERIC Document Reproduction Service No. EJ 410 730). Kohen, E., 1974. A validity study for the CLEP introductory calculus subject examination at the University of Illinois. Research Report #349. (ERIC Document Reproduction Service No. ED 087 295). Lamar University: 1923 - Present History. Beaumont, Texas: Lamar University. Retrieved May 30, 2002 from the World Wide Web: http://www.lamar.edu/admin/history/index.html McLure, G. T., A. Sun, M. J. Valiga. 1997. Trends in advanced mathematics and science course taking and achievement among ACT-tested high school students: 1987-1996. (ERIC Document Reproduction Service No. ED 417 966), 68 pp. Noble, J. P. & R. Sawyer. 1988. Predicting grades in specific college freshman course from ACT test scores and self-reported high school grades. (ERIC Document Reproduction Service No. ED 322 162), 61 pp. O’Heam, M. 1986. The ACT assessment: implications of recent research. (ERIC Document Reproduction Service No. ED 250 330), 23 pp. Peng, C. Y. J. & T. S. H. So. 2002. Logistic regression analysis and reporting: a primer. Understanding statistics, 1,1, 31-70. Reiner, J. R. 1971. Differential effects of initial course placement as a function of ACT mathematics scores and high school rank-in-class in predicting general performance in chemistry. Educational and psychological measurement, 31, 4, 977-81 (ERIC Document Reproduction Service No. EJ 050 356), 5 pp. Sweet, S. A. & K. Grace-Martin. 2003. Data analysis with SPSS a first course in applied statistics. Allyn and Bacon (2"^ Edition), 236 pp. UTSA 2000 Highlights: UTSA - An Overview. San Antonio, Texas: The University of Texas at San Antonio. Retrieved May 29, 2002 from the World Wide Web: http://www.utsa.edu/hilights/overview.htm PFM at: messina@uiwtx.edu TEXAS J. SCI. 60(3):201-214 AUGUST, 2008 ALGAL COMMUNITY STRUCTURE OF THE EAST AND WEST FLOWER GARDENS, NORTHWESTERN GULF OF MEXICO Erin M. Hill and Roy L. Lehman Center for Coastal Studies, Texas A&M University-Corpus Christi 6300 Ocean Drive, Corpus Christi, Texas 78412 Abstract.-The Flower Garden Banks are located approximately 200 km south of the border between Texas and Louisiana, on the outer continental shelf, beginning at a depth of approximately 20 m. The algal community has not been comprehensively evaluated and only checklists of a few dominant macroalgal species have been reported. This study utilized both destructive and non¬ destructive techniques to characterize the algal community structure. Harvest and photogrammetric samples were collected during two trips to the Flower Garden Banks in October 1998 and March 1999. A total of forty 0.25 m“ quadrats of standing stock material were randomly collected, curated and stored for further evaluation. Photogrammetric samples (161) were obtained using an underwater camera and were evaluated by a grid of 25 random points. A total of 4,025 points of information were used to calculate percent composition and cover. Harvest sampling was used to characterize the algal composition of the ‘red algal mat’ which was the dominant feature comprising 38.4% of the photogrammetric samples. A total of 44 species were identified from the samples. The ‘red algal maf was primarily composed of members of the Order Ceramiales {Centroceras, Ceramium, Hypoglossiim, Polysiphonia and Anotrichium). The Flower Garden Banks are deep-water coral habitats located on the edge of the outer continental shelf in the northwestern Gulf of Mexico and cover an area of 143 km“. The Flower Garden Banks are found at approximately 18-36 m below the water surface and extend down to 100-150 m (Lugo- Femandez et al. 2001). The Flower Garden Banks are separated into the East Flower Gardens and the West Flower Gardens and are 12-15 km apart. They were designated as a protected National Marine Sanctuary in 1992. The East Flower Garden Bank is located at 27° 54.5' N latitude and 93° 36.0' W longitude, approximately 193 km southeast of Galveston, Texas. The West Flower Garden Bank is located approximately 172 km southeast 202 THE TEXAS JOURNAL OF SCIENCE-VOL.60, NO. 3, 2008 of Galveston, Texas at 27° 52.4' N latitude and 93° 48.8' W longitude (Fig. 1). Both banks are topographic features created by the uplift of underlying salt domes of Jurassic, Louann origin (Rezak 1981). Bedrock material above the domes is covered with an overgrowth of calcareous marine organisms and represents the largest charted calcareous banks in the northwestern Gulf of Mexico (Bright et al. 1985) and the northernmost coral reefs on the continental shelf of North America (Bright et al. 1984). The Flower Garden Banks are part of a widely dispersed and discontinuous arc of reef material located along the outer continental shelf of the Gulf of Mexico (Rezak et al. 1985). The diversity of the coral community at the Flower Garden Banks has been described as depauperate as it supports only 20 species of hermatypic corals (Bright et al. 1984; Lugo-Femandez et al. 2001). Although low diversity communities exist on neighboring banks, the reefs adjacent to Cabo Rojo, approximately 100 km south of Tampico, Mexico are the closest extensively developed coral reefs in the Gulf of Mexico and make up the Tuxpan Reef System (Universidad Veracruzana 1996; 2003; Tunnell 2007). The Tuxpan Reefs are approximately 644 km south of the Flower Garden Banks and are composed primarily of stony corals; 31 species of scleractinia coral and two hydrocorals (Universidad Veracruzana 1996; 2003; Tunnell 2007). Monitoring of the Flower Garden Banks began in 1989 (Gittings et al. 1992) for the purposes of: (1) documenting long¬ term natural changes in reef growth and associated communities, (2) providing information and scientific data on the effect of gas and oil exploration in the vicinity of this sensitive ecosystem, and (3) encouraging and coordinating research efforts with agencies and institutions in order to better evaluate and/or determine causes of environmental change in the vicinity of the Flower Garden Banks (Dokken et al. 2001). Taxonomic lists of benthic HILL & LEHMAN 203 Figure 1. Map showing location of East and West Flower Garden Banks. marine algae from the Flower Garden Banks were included in investigations by Bright and Pequegnat (1974) and Rezak et al. (1985; 1990). This study characterizes and provides an updated taxonomic assessment of the algae, coral and algae associations and algal distribution of the East and West Flower Garden Banks. Materials and Methods Ecological field methods followed those described by Littler & Littler (1985) and Brower et al. (1989). A herbarium voucher collection of benthic macroalgae (seaweeds) was completed and is located at the Ruth O’Brien Herbarium at Texas A&M University-Corpus Christi, Corpus Christi, Texas. Voucher specimens of macroalgal samples were also preserved in 2% gluteraldehyde and microscopic material was preserved on microscope slides sealed with Flo-Texx. 204 THE TEXAS JOURNAL OF SCIENCE-VOL.60, NO. 3, 2008 Photogrammetric and harvest samples were collected during two trips to the Flower Garden Banks in October 1998 and March 1999. Random quadrats were photographed using a Nikonos V equipped with a 28 mm lens, close-up kit, and Nikonos strobe. Each transparency represented a 0.25 m^ quadrat that was evaluated by projecting onto a grid containing 25 random points (modified point quadrat) and evaluated based on (1) Bare Substrate, (2) Cyanobacteria, (3) Green Algal Mat, (4) Red Algal Mat, (5) Coralline Algae, or (6) Live Coral coverage. Values are expressed as the number of ‘hits’ for each species or feature divided by the total number possible. This number varied with each bank and season measured. A total of 4,025 points of data were identified and used to calculate percent composition, species richness, dominance, diversity and cover. Such samples can be used to generate precisely detailed and highly reproducible quantitative information, including cover, density, frequency and diversity (Littler & Littler 1985). Diversity and other indices were calculated from relative abundance of both, algae and total species observations. Diversity values were determined for each bank and sampling season using the Shannon Index (H', natural log) (Shannon 1948), Richness, Evenness (T) (Pielou 1966) and Simpson’s Index of Dominance (Simpson 1948). Relative Community Similarity was determined by species or feature abundance from each bank and season and compared with total percent similarity and Morisita’s Index of Community Similarity (Morisita 1959). Morisita’s Index is based on Simpson’s Index of Dominance and determines the probability that two randomly selected individuals will be the same species. The range of results are 0 (no similarity) to approximately 1 (identical). Macroalgal mat samples were harvested using paint scrapers and placed into nylon net bags. A total of forty 0.25 m^ quadrats of standing stock material were randomly collected, preserved HILL & LEHMAN 205 and stored for evaluation. This sampling was used to characterize the algal composition of the ‘red algal mat’. Results A total of forty 0.25 m quadrats of standing stock material were randomly collected along with one hundred sixty-one 0.25 m^ photo-quadrats from an average depth of 27 m. A systematic list of marine algae from the East and West Flower Garden Banks has been compiled (Table 1). The systematic organization follows that of Wynne (1998). Taylor (1960) and Schneider & Searles (1991) were referenced when species were encountered that were uncommon to the region. Photogrammetric sampling -ThQ feature ‘red algal mat’ had the greatest coverage comprising 38.4% of all photogrammetric samples followed by Bare Substrate (14.6%), Coralline Algae (9%), Green Algal Mat (8.1%), and Cyanobacteria (4%) (Figure 2). Species richness totaled 25 with both banks combined. The East Bank had the greatest species richness October 1998 with 19 and lowest March 1999 with 14 (Table 2). Diversity measures using Shannon’s Diversity Index and Evenness show little differences in diversity between years and between the East and West Banks (Table 2). The West Bank had the greatest diversity October 1998 (H' = 2.10) and the East Bank had the highest diversity March 1999 (H' = 2.06). Overall, species diversity with both banks combined was H' = 2.20. Morisita’s Index of Community Similarity indicated the East and West Banks in 1999 and the total of the East and West Banks from 1998 & 1999 were almost identical (0.94). Percent (Proportional) similarity supported these results (Table 3). Association between corals and algal mat communities were investigated to determine if a particular coral species had a partiality with the red algal mat. Within all quadrats, each time a coral species was encountered, the presence or absence of the red 206 THE TEXAS JOURNAL OF SCIENCE-VOL.60, NO. 3, 2008 Table 1. Taxonomic list of algae collected from East and West Flower Garden Banks, northwestern Gulf of Mexico. (E=East, W=West). DIVISION CYANOBACTERIA Order Oscillatoriales Family Oscillatoriaceae Lyngbya sp. E& W Oscillatoria sp. E& W Schizothrix calcicola (C. Agardh) Gomont E& W Spirulina sp. W Unknown (3 species) E& W DIVISION RHODOPHYTA Order Compsopogonales Family Erythropeltidaceae Erythrocladia sp. E Erythrotrichia sp. E Order Bonnemaisoniales Family Bonnemaisoniaceae Asparagopsis taxiformis (Delile) Trevisan E & W Order Corallinales Family Corallinaceae Amphiroa fragilissima{L\rm2iQm) Lamouroux E Corrallina spp. (3 species) E & W Jania adhaerens Lamouroux E & W Order Ceramiales Family Ceramiaceae Anotrichiiim temie (C. Agardh) Nageli E & W Antithamnion sp. E & W AntithamnioneUa sp. E CaUithamniella sp. E & W Callithanmion sp. E & W Centroceras clavidatum (C. Agardh) Montagne E & W Ceramhim spp. (2 species) E & W Griffithsia globulifera Kiitzing W Plenosporium flexuosum (C. Agardh) De Toni E & W Ptilothamnion occidentale Searles W Rhododictyon bermudeme W. R. Taylor E Spyridia spp. (2 species) E & W Family Delesseriaceae Hypoglosswn hypoglossoides (Stackhouse) Collins et Hervey E & W Hypoglossum sp. W Myriogf^amme sp. W E& W Family Dasyaceae Dasya sp. E & W HILL & LEHMAN 207 Table 1 (Cent.) Family Rhodomelaceae Chondria sp. E& W Herposiphonia sp. W Polysiphonia spp. (3 species) E& W Order Rhodymeniales Family Rhodymeniaceae Botryocladia occidentalis (B0rgesen) Kylin E& W Chrysymenia enteromorpha Harvey E& W Chrysymenia sp. W Rhodymenia divaricata Dawson E Order Gigartinales Family Dumontiaceae Dudresnaya sp. W Family Hypneaceae Hypnea sp. W Order Gelidiales Family Belidiaceae Gelidium sp. w DIVISION OCHROPHYTA Class Phaeophyceae Order Dictyotales Family Dictyotaceae Dictyota dichotoma (Hudson) Lamouroux E& W Dictyota sp. E& W Dictyota sp. E& W Lobophora variegata (Lamouroux) Womersley E& W Spatoglossum schroederi (C. Agardh) Kiitzing E& W DIVISION CHLOROPHYTA Order Ulotrichales Family Ulotrichaceae Ulothrix flacca (Dilwyn) Thuret E& W Order Dasycladales Family Polyphysaceae Polyphysa polyphysoides (P. and H. Crouan in Maze and Schramm) Schnetter E& W Order Ulvales Family Ulvellaceae Entocladia viridis Reinke E Family Ulvaceae Enteromorpha sp. E& W Order Caulerpales Family Bryopsidaceae Bryopsis pennata Lamouroux E& W Derbesia sp. E& W 208 THE TEXAS JOURNAL OF SCIENCE-VOL.60, NO. 3, 2008 Table 1 (Cont.) Family Caulerpaceae Caulerpa sp. W Order Cladophorales Family Cladophoraceae Chaetomorpha sp. E Cladophora sp. E & W Family Anadyomenaceae Anadyomene stellata (Wulfen) C. Agardh W Bare Cyanobacteria Coralline Green Mat Red Mat Substrate Algae □ TD 98 STD 99 SFG TD % Figure 2. Percent algal coverage of photogrammetric samples from 1998, 1999, and both years combined. (TD = total; FG = Flower Gardens) algal mat was assessed. A proportion of coral species to total red algal mat was computed. Montastrea cavernosa was most often associated with the red algal mat at 54.2% with Diploria strigosa following at 27.1%. Additional corals that were also associated with the red algal mat were Diplora clivosa at 11.9% and Montastrea annularis at 10.1%. Harvest sampling -WdiVVQsi samples were evaluated to characterize the species composition of the algal mat found at the coral/algal interface. The most common species collected were fflLL & LEHMAN 209 Table 2. Species richness, diversity, dominance and evenness of the East and West Flower Gardens Banks, 1998-1999. Bank/ Year Species Richness Shannon’s Diversity Shannon’s Evenness Simpson’s Dominance Simpson’s Evenness West 98 15 2.10 0.77 0.15 0.93 East 98 19 1.90 0.65 0.26 0.77 West 99 16 1.94 0.70 0.23 0.82 East 99 14 2.06 0.78 0.18 0.88 Total 25 2.20 0.68 0.19 0.84 Table 3. Community similarity using Percent Similarity and Mortisita’s Index of the East and West Flower Garden Banks, 1998 & 1999. BankAfears Percent Mortisita’s Similarity Index East 98 vs. West 98 53.01 0.72 East 99 vs. West 99 72.26 0.94 East 98 vs. East 99 61.54 0.85 West 98 vs. West 99 58.94 0.79 East vs. West Total 73.59 0.94 from the Division Rhodophyta. A ‘red algal mat’ was the dominant algal and non-algal coverage comprising 38.4% of all photogrammetric samples. The mat was composed primarily of members of the Order Ceramiales comprising 37.6% of coverage and was represented by species of Centroceras, Ceramium, and Polysiphonia (Table 3). The Order Ceramiales was also the dominant group at each bank; 38.3% at the East, and 36.9% at the West. The East bank was represented by 13 algal families and the West Bank by 14. A total of 44 species were identified from harvested samples collected from the East and West Flower Gardens Banks (Table 4). Bryopsis pennata and Derbesia, both in the Order Caulerpales, composed 12.0% of the ‘green algal mat’ on the East Flower Gardens and 11.0% on the West Flower Gardens. The Rhodophyta, Jania adhaerens, was the most common 210 THE TEXAS JOURNAL OF SCIENCE-VOL.60, NO. 3, 2008 Table 4. Total percentage of each algal species from the East and West Flower Garden Banks 1998-1999 contributing to total coverage. Species East Bank West Bank Amphiroa fragilissima 3.4 0 Anotiichium tenue 0 2.3 AntithanmioneUa sp. 1.5 0.4 Asparagopsis taxiformis 4.1 2.7 Botryocladia occidentalis 1.1 1.1 Bryopsis pennata 6 6.5 CaUithamnion sp. 1.5 2.3 Caiilerpa sp. 0 1.1 Centroceras clavidatum 7.5 2.7 Ceram him sp. 6.4 3.8 Chaetomorpha sp. 0.8 0 Chondria sp. 3.8 2.7 Chrysymenia enteromorpha 1.5 2.3 Cladophora sp. 4.9 3.4 Coral Ha a sp. 0.4 0 Dasya sp. 1.1 1.5 Derbesia sp. 6 3.4 Diatom mat 0 0.4 Dictyopteris membranaceae 0 1.1 Dictyota sp. 3.8 3 Dudresaaya sp. 0 1.9 Enterocladaia virdis 0.8 0 Enteromorpha sp. 0.8 2.7 Eiythrocladia sp. 0.4 0 Erythrotrichia sp. 1.5 0 Gelidhim sp. 0 1.1 Griffithsia globulifera 0 2.7 Herposiphonia sp. 0 1.5 Hypnea sp, 0 0.8 Hypoglossum hypoglosoides 1.9 2.7 Jania adhaerens 7.1 6.1 Lobophora variegata 4.5 1.5 Lyngbya sp. 7.5 5.7 Myriogramme sp. 0 0.4 Oscillatoria sp. 0 6.8 Plenosporiiim flexuosum 3 3.8 Polyphysa polyphysoides 1.5 0 Polysiphonia sp. 4.9 5.3 Ptilothamnion occidentale 0 0.4 Rhodymenia divaricata 0.8 0 Spatoglossum schroederi 1.9 0.4 Spirulina sp. 0 0.8 Spyridia sp. 3.4 4.9 Ulothrix flacca 2.6 3.4 Unknown 3.8 6.5 HILL & LEHMAN 211 coralline alga found at both Banks with coverage of 6.1% at the West and 7.1% at the East Bank. Cyanobacteria; primarily Oscillatoriales, were common in samples, representing 7.5% at the East Flower Gardens, and 11.0% at the West Flower Gardens, Oscillatoriales was represented by only a few taxa with Lyngbya as the most common genus at both banks. Discussion It is difficult to determine causes or effects of algal populations affecting the health of particular coral species of the Flower Garden Banks because the limited opportunity of sampling during this study. When given the opportunity to collect algal material from the Flower Garden Banks, especially at the interface of coral, algal material should be photographed in situ and samples collected, preserved in 2% gluteraldehyde, and returned to the laboratory for identification and evaluation. Coral species, depth, temperature and other environmental factors at the site should be recorded. Ceramiales is an Order composed mainly of annual species that dominate as opportunists at sites with irregularly fluctuating conditions (i.e., temperature) (Luning 1990). These opportunists are quick growing algae that, after bacteria and diatoms, are the first multicellular algae to appear on bare, nonliving or damaged substrates. These algae exhibit variability in biomass depending upon herbivorous grazing or changes in environmental and seasonal conditions, Guimaraens & Coutinho (1996) evaluated the effect of an upwelling region near Rio de Janeiro, Brazil on the spatial and temporal variation of benthic marine algae. The survey of benthic algae yielded groups including the Ceramiales with temperature affinities occurring in sites directly influenced by upwelling waters. Their data showed water temperature affects distribution and abundance of algae enabling warm 212 THE TEXAS JOURNAL OF SCIENCE-VOL.60, NO. 3, 2008 temperate species (18”20° C) to survive under otherwise tropical conditions. A seven year mean of water temperature recorded at a depth of 24 m at the West Flower Garden Banks resulted in a range of 18- 30®C with low temperatures occurring in February (18-20®C) and highs in July- August (29-30°C) (Lugo-Femandez 1998). This temperature range at the Flower Garden Banks is optimum for the growth of the ‘red algal mat’ composed primarily of members from the Order Ceramiales (Guimaraens & Coutinho 1996). The increased growth of these algae may become an environmental problem, possibly global, that would be practically impossible to mediate. The best action is to monitor changes over time to determine the magnitude and effect to the reef resulting from competition of the ‘red algal mat’. Little is known about the community dynamics of benthic marine algae and their effect on the character of biotic reefs. Information on the role of inter¬ specific and intra-specific competition, recruitment, natality and mortality phenomena is lacking and it is difficult to make definitive statements concerning the role of algae and their effect on other organisms. Acknowledgments Thanks are extended to the Flower Garden Banks National Marine Sanctuary for their support, interest and permission to conduct this ancillary project in conjunction with the MMS/NOAA funded project “Long-term monitoring at the East and West Flower Garden Banks, 1998-1999.” Thanks to Dr. Quenton Dokken, Dr. Carl Beaver, Susan Childs and Suzanne Bates, Center for Coastal Studies, Texas A&M University- Corpus Christi for their support in this project. We appreciate Lee Lehman for his help in sample collection and Amy Nunez for HILL & LEHMAN 213 coral identification, Texas A&M University-Corpus Christi, and the crew of the M/Y Spree for their assistance. Literature Cited Brower, J. E., J. Zar & C. N vonEnde. 1997. Field and laboratory methods for general ecology, WCB/McGraw-Hill, St. Louis, Missouri, 273 pp. Guimaraens, M. & R. Coutinho. 1996. Spatial and temporal variation of benthic marine algae at the Cabo Frio upwelling region, Rio de Janeiro, Brazil. Aquat. Bot., 52:283-299. Littler D. S., M. M. Littler, K. E. Bucher & J. N. Norris. 1989. Marine plants of the Caribbean: A field guide from Florida to Brazil. Smithsonian Institution Press, Washington, D.C., 249 pp. Lugo-Femandez, A. 1998. Ecological implications of hydrogeography and circulation to the Flower Gardens and Stetson Banks, northwest Gulf of Mexico. Gulf Mex. Sci., 2:144-160. Lugo-Femandez, A., K. J. P. Deslarzes, J. M. Price, G. S. Boland & M. V. Morin. 2001. Inferring probable dispersal of Flower Garden Banks coral larvae (Gulf of Mexico) using observed and simulated drifter trajectories. Cont. Shelf Res., 21:47-67. Liining, K. 1990. Seaweeds. Their environment, biogeography, and ecophysiology. John Wily & Sons, Inc., New York, 527 pp. Morisita, M. 1959. Measuring of interspecific association and similarity between communities. Mem. Fac. Sci. Kyushu Univ., Ser. E (Biol.), 3:65-80. Pielou, E. C. 1966. The measurement of diversity in different types of biological collections. J. Theoret. Biol., 13:131-144. Schneider, C. W. & R. B. Searles. 1991. Seaweeds of the Southeast United States, Cape Hatteras to Cape Canaveral. Duke University Press, Durham, NC, 553 pp. Shannon, C. E. 1948. A mathematical theory of communication. Bell System Tech. J., 27:379-423, 623-656. Simpson, E. H. 1949. Measurement of diversity. Nature, 163:688. Taylor, W. R. 1960 Marine algae of the eastern tropical and subtropical coasts of the Americas. University of Michigan Press, Ann Arbor, MI, 870 pp. Tunnell, J. W., Jr. 2007. Reef distribution (chapter 2). In Tunnell, J.W. Jr., Chavez, E.O. and Withers, K. (eds) Coral reefs of the southern Gulf of Mexico. Texas A&M University Press, College Station, TX, 216 pp. Universidad Veracmzana. 2003. Documento tecnico justificativo para la creacion de un area natural protegida en el sistema arrecifal Lobos-Tuxpan. University Press, College Station, TX, 216 pp. Universidad Veracmzana. 1996. Ecobuceo en la Costa Norveracruzana. Facultad de ciencias biologicas y agropecuarias grupo de ecologia arrecifal. 214 THE TEXAS JOURNAL OF SCIENCE-VOL.60, NO. 3, 2008 Wynne, M. J. 1998. A checklist of benthic marine algae of the tropical and subtropical western Atlantic: First Revision. Gebruder Bomtraeger. Berlin, Germany, 155 pp. EMH at: Erin.Hill@tamucc.edu AUGUST, 2008 TEXAS T OF SCI. 60(3):2 15-220 A COMPARISON OF MANDIBULAR STRUCTURE IN RUDDY AND MASKED DUCKS {OXYURA JAMAICENSIS AND NOMONYX DOMINICU^ Jack C. Eitniear and Kent Rylander 218 Conway Drive San Antonio, Texas 78209 and Texas Tech University-Junction Center P.O. Box 186, Junction, Texas 76849 Abstract.-In order to test inferred differences in foraging behavior of both the masked duck {Nomonyx dominicus) and the ruddy duck (Oxyura jamaicensis), this study examined nine characters and three ratios in the mandibles of 25 male and 25 female museum specimens of both species. Statistical analysis indicated that O. jamaicensis had longer lamellae and a slightly broader bill than N. dominicus (data for both sexes combined), which may adapt O. jamaicensis for efficiently sifting through bottom silt. Specimens of A dominicus exhibited a stouter bill (greater height/length ratio of the maxilla), supporting the premise that this species is principally adapted for grazing on tough plants. Most bill characters were greater in length or width in O. jamaicensis as opposed to the masked duck and in ruddy duck males as opposed to ruddy duck females. Also, maxilla height (absolute as well as relative to dorsal maxilla length) was greater in male than in female masked ducks, suggesting a possible signal function in bill shape. Based on bill morphology, this study supports the premise that the genus Nomonyx is less advanced than the Oxyura/Biziura group. Ducks of the tribe Oxyurini are distinguished from all other waterfowl by their elongated and pointed tail feathers and stiffened shafts (Johnsguard & Carbonell 1996). Within the tribe Livezy (1995) proposed the generic addition of Nomonyx (originally proposed by Salvador! 1895) differentiating the masked duck {Nomonyx dominicus) from other Oxyura based on a variety of characters including morphology and behavioral traits. Considering the premise that morphology represents aspects of the relationships between the organism and its environment (Bock 1977), this study examined mandibular traits of masked and ruddy ducks as a reflection of their foraging ecology. Methods and Materials The following measurements were made on 50 museum specimens (25 male and 25 female) of both Nomonyx dominicus and Oxyura jamaicensis: length of mandible, length of maxilla 216 THE TEXAS JOURNAL OF SCIENCE- VOL. 60, NO. 3, 2008 (dorsal), length of maxilla (lateral), depth of maxilla, width of maxilla, width of mandible, length of tarsus (as an indicator of general body size), and number and size of lamellae (Table 1). Lamellar height was measured in the most rostral 30 lamellae. Using a low-power dissecting microscope, flattened insect pin tips that were calibrated to 0.4, 0.6, 0.8, 1.0 mm (with a stage micrometer) were inserted between adjacent lamellae (Fig. 1). All other measurements were made with dial calipers. Due to the difficulty in many specimens of exposing the most caudal lamellae, the authors estimate an accounting error of some 4-8 lamellae. Descriptive statistics and student's ^test were done with Statview (1996) version 4.51 for Macintosh. Material examined museum specimens (25 male and 25 female) of Oxyura jamaicensis from nine countries and fifty specimens (25 male and 25 female) of Nomonyx dominicus from 12 countries were examined during this study. Accession numbers of specimens are from the following museums: Museum of Vertebrate Zoology, University of California, Berkeley (MVZ); Texas Cooperative Wildlife Collection-College Station (TCWC), Field Museum of Natural Flistory-Chicago (FMNH), Louisiana Natural History Museum-Baton Rouge (LSUMZ) and Texas Tech University Museum-Lubbock (TTU). Oxyura jamaicensis -USA (LSUMZ 6791, 10406, 21507, 25271, 39136, 43115, 43116, 80214, 154354, 155831, 157507, TTU 178, 618, 3076, 4012); MEXICO (LSUMZ 10697, 14900, 27127, 43118); PUERTO RICO (LSUMZ 10336, 23185); BAHAMAS (LSUMZ 141425, 141426, 141427); DOMINICAN REPUBLIC (LSUMZ 141430, 141431, 141432, 141433); HAITI (LSUMZ 141428, 141429); EL SALVADOR (LSUMZ 50481); PERU (ESUMZ 34550, 34551, 63848); BOEIVIA (LSUMZ 35633, 35634, 35635, 35636, 35637, 37023, 37024, 37025, 37026, 37027, 37028, 37029, 37030, 37031, 37032, 37033). Nomonyx dominicus. -US A (FMNH 96714, TCWC 11150); JAMAICA (FMNH 96719); CUBA (FMNH 96717, 96718, 156703, 411026, 411028, 411029); COSTA RICA (FMNH 126953, 127846, EITNIEAR & RYLANDER 2 S D g !+ ^ E" D o r« « 3 TO C. K> p KJ P p p o a\ L-* E la b b bs ho » O 0n» b b b b b b b ho UJ o Ul m UJ © Ki o © KJ © © Oh O o Ohi o .Su © fv.> w w K> p o o p o “ o — • p p P ►=“»* P K) p OJ Ni O o o Ud o b ka b k> e© b Ln b b b b b b© b b b b b UJ o o© a© oo 4i. o O tv3 o o o© o -a oo Oh *— SO Oh ^4 so K) W O ^ ©O to O O O O O p O o O 4a. K4 04 04 ►— lO p p p 4^ m p «b U> p p b Oh b b b b b b on b b b b b b w b 04 ©S ©« b b u> tsj on o© o o o O 04 © OA © © © © SO O OOOOOj-^M**. tsj — 4i^ UJ 0^»— — 0N^4^00 oLno'-fco’-^OJOsisfc^^soLAE^EoobslAbowoEP-^ W-^W^fsJO-4O^Os0O©©O^OU»O'-4OU^O<-rt©© 4SkfO— 0NOpp;^ppsO0\;~*Wi~.\0OO lo^EKjlaEbo'-^P-^i^boooKioPji^oE^ 00©orsJ04s.O'^0'--aO'«DOOsOsOO<-nON opopp^^p^ppooppp^ K>wu»->4000 0‘>40 OWOsOO K> O •— to k> ^ o o o r* P w ©e O Cu Cu v< ?3 c Cu o. c a. 217 Table I . Measurements of bill dimensions in ruddy and masked ducks. Numbers are means (mm). The greater value is listed in bold-faced type when the probability is less than 0.05 that the two samples are from the same population. 218 THE TEXAS JOURNAL OF SCIENCE- VOL. 60, NO. 3, 2008 Figure L Ventral view of ruddy duck (left) and masked duck (right) to show lamellae (arrow). Lamellar length was measured by inserting calibrated pins in the spaces between the lamellae. 127847, 156705, 156706, 156707, 156708, 408630, 408631, 409925); PANAMA (FMNH 409924); COLOMBIA (FMNH 13611, 416473, 418705, 422612; MVZ 140619); VENEZUELA (FMNH 400007, 400008); GUYANA (FMNH 32190, 32191); SURINAME (MVZ 123495); ECUADOR (FMNH 414321); PARAGUAY (FMNH 411008, 411212, 412533, 412534, 413272, 413273, 413274, 413275, 413276, 414416, 416533, 416534, 416536, 417165; MVZ 99594, 99595, 99596); BRAZIL (FMNH 410472, 411414). Results and Discussion Table 1 shows that both species are sexually dimorphic with regard to bill size, and that the density of lamellae is approximately the same in both species, suggesting that if the ruddy duck is better adapted for sifting through silt, the adaptive advantage relates to lamellar size rather than number. On the other hand, the difference in scale between lamellar size and body size is so great that the larger lamellae may simply be a function of the overall larger size of the ruddy duck. The relative width of the bill is slightly larger in the ruddy duck: the ratios of maxilla width to dorsal maxilla length are 0.57 (ruddy duck) EITNIEAR & RYLANDER 219 and 0.52 (masked duck), both sexes combined. Relative bill height (maxilla height/dorsal maxilla length) is greater in the masked duck (0.49) than in the ruddy duck (0.45), sexes combined, suggesting that the masked duck possesses a stouter bill. Where morphological and ecological measurements have been compared morphological generally predicts a greater portion of the variation in ecology than ecology does of morphology (Wainwright & Reilly 1994). Field data documents that adult breeding ruddy ducks feed almost exclusively on invertebrates by inserting their bills into the substrate, and moving the head in a lateral arc about 1.5 times the width of the bill, forcing the substrate out of the bill through the lamellae (Johnsgard & Carbonell 1996). During the non-breeding season ruddy ducks feed on a more varied diet that includes vegetative materials (Brua 2001). While little information is available on masked ducks specimens collected indicate that this species feeds primarily on seeds, stems, leaves and roots of aquatic plants. Nesting adults may feed on aquatic insects and crustaceans, but few data are available (Eitniear 1999). The literature on masked ducks also documents its preference for bodies of water with dense emergent vegetation often residing within the vegetation itself The ruddy duck, however, prefers open areas of shallow water (Anderson & Tacha 1999; Eitniear & Colon 2005). While the masked duck’s bill is high and broad at the base, as in other stifftails, it is not as depressed at the tip; nail is less narrow and recurved, suggesting that perhaps roots and stems are a key part of the diet (Johnsgard & Carbonell 1996). The ruddy duck’s longer lamellae and slightly broader bill may adapt this species for efficiently sifting through bottom silt. The resulting analysis of this study supports the premise that the masked duck is more primitive in structure being less adapted for sifting but retaining a stout mandible whose anatomy facilitates aquatic plant extraction. Ecology and morphology provide alternative, but mutually consistent expressions of the outcome of ecological and evolutionary adjustments between phenotype and environment (Bock 1977). Based on bill morphology, this study supports the premise that the genus Nomonyx is less advanced that the Oxyura/Biziura group 220 THE TEXAS JOURNAL OF SCIENCE- VOL. 60, NO. 3, 2008 (Johnsgard & Carbonell 1996). Additional field and captive studies should provide insights as to whether morphology reflects feeding ecology or if mandibular morphology is less important than behavioral foraging strategies (Guillemain et al. 2002) in this cryptic, elusive species of stifftail duck. Acknowledgments We are grateful to the following persons for the institutional loans of study skins: K. Arnold, S. Cardiff, C. Cicero, N. McIntyre, K. Tsao, J. Van Remsen and D. Willard. Appreciation is extended to J. T. Baccus and M. Small for their comments on an earlier draft of the manuscript. Literature Cited Anderson, J. A & T. C. Tacha. 1999. Habitat use by Masked Ducks along the Texas Gulf Coast. Wilson Bulletin 111:119-121. Bock, W. J. 1977. Toward an ecological morphology. Die Vogelwarte 29:127-135. Brua, R. B. 2001. Ruddy Ducks {Oxyura jamaicensis) in The Birds Of North America, No. 696. (A. Poole and F. Gilfeds.). The Birds of North America, Inc., Philadelphia, 46 pp. Eitniear, J. C. 1999. Masked Duck {Nomonyx dominica) in The Birds of North America, No. 393 (A. Poole and F. Gill, eds.). The Birds of North America, Inc. Philadelphia, 12 pp. Eitniear, J. C. & S. Colon. 2005 Recent observations of Masked Duck Nomonyx dominica in the Caribbean. Caribbean Jour. Science 41(4):861-864. Guillemain, M., H. Fritz, N. Guillon & G. Simon. 2002. Ecomorphology and coexistence in dabbling ducks: the role of lamellar density and body length in winter. Oikos 98:547-551. Johnsgard, P. A. & M. Carbonell. 1996. Ruddy Ducks & other stifftails. Univ. of Oklahoma Press, Norman, 291 pp. Livezy, B. C. 1995. Phylogeny and comparative ecology of stiff-tailed ducks (Anatidae:Oxyurini). Wilson Bulletin, 107:214-234. Salvador!, T. 1895. Catalogue of the Chenomorphae (Palamedeae, Phoenicopteri, Anseres), Crypturi and Ratitae in the collection of the British Museum, Longmans, London, England. STATVIEW. 1996. Version 4.51 for Macintosh. Abbacus Concepts, Inc., Berkeley, CA. Wainwright, P. C. & S. M. Reilly. 1994. Ecological Morphology. Univ. of Chicago Press, Chicago, Illinois, 367 pp. JCE at: admin@cstbinc.org TEXAS J. SCI. 60(3), AUGUST, 2008 221 GENERAL NOTES THE OCCURRENCE OF AGARDHIELLA RAMOSISSIMA (GIGARTINALES) AND ACANTHOPHORA SPICIFERA (CERAMIALES) IN THE TEXAS COASTAL BEND Ryan L* Fikes and Roy L. Lehman Center for Coastal Studies, Texas A&M University-Corpus Christi 6300 Ocean Drive, Corpus Christi, Texas 78412 The algal flora of the Texas coast is largely of tropical affinity; however, a distinctive cool temperature element develops during the winter and early spring (Edwards & Kapraun 1973). This range in water temperatures gives the Texas coast a variation of macroalgae characteristic of both surrounding areas and the tropics. Studies along the Texas coastline over the past few decades have contributed to new taxa for those areas (e.g.Kapraun 1980, Medlin 1984, Wardle 1992). The Corpus Christi Bay area is now known to support at least 112 species of algae (Lehman 1999). Within this list, 24 additional species of macroalgae are reported that had not been previously noted in the Corpus Christi Bay area by past research, nine of which have a tropical affinity. A study was recently completed of the colonization of macroalgae along the rocky jetties of Packery Channel (27° 36.836' N; 97° 12.044' W), a new man-made tidal inlet opened in 2006 near Corpus Christi, Texas. Macroalgae were sampled during bimonthly destructive sampling from September 2006 thru March 2007. Specimens were obtained by scraping clear quadrats (20 X 30 cm) into 500 pm biobags at a depth of < 1 m. By the end of the year-long sampling event, 40 species of macroalgae had been collected from the inlet. 222 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 This study reports the occurrence of Agardhiella ramosissima (Harv.) Kylin, a tropical species found occurring on coral reefs. In the Atlantic, this species has been reported from as far north as North Carolina (Schneider and Searles 1973; 1991) and south as Rio de Janeiro, Brazil (Pedrini 1984), but has never been reported from the northwest Gulf of Mexico. Less than 20 individual plants were collected, with size ranging from 1-5 cm. Specimens were further confirmed by the presence of zonate tetraspores embedded in surface patches within the thalli. Dry-pressed specimens are housed with the Phycological Collections of the Center for Coastal Studies Herbarium of Texas A&M University- Corpus Christi (accession number PCK 019 ramosissima). Cheney’s Floristic Ratio (Cheney 1977) was used to indicate the floristic type represented by the algae growing at Packery Channel and was found to be 9.0 for this study. This is exceptionally high when compared to other macroalgal communities from the Gulf Coast. Temperature is the major factor controlling geographical distribution of marine algae (Edwards and Kapraun 1973) and, therefore, high ratios (meaning a more tropical flora) should be found progressing from north to south. This finding for Packery Channel is likely to be inaccurate due to incomplete development of the community. The authors are of the opinion that Cheney’s Floristic Ratio, while useful in classifying communities, should be avoided in developing habitats. Jetties are likely to draw at least part of their populations from the floating pool of individuals in the Gulf of Mexico (Britton & Morton 1989). Until jetties were constructed along the Texas coast, our outer shores were limited in algal growth, for they generally lack the necessary hard substrate. The inhibition model for structural assemblages states that any species can potentially colonize bare space (Connell & Slatyer 1977). With increased human impact in coastal zones and the increase in numbers of TEXAS J. SCI. 60(3), AUGUST, 2008 223 jetties and seawalls, both the algal diversity and documentation of this change have increased significantly. In a related study conducted simultaneous to that along the Packery Channel jetties, Acanthophora spicifera (Vahl) Borgesen was identified from the recessed portion of the channel. Several (10-15) specimens of this species were collected near the intersection of the Laguna Madre and the Gulf Intracoastal Waterway (GIWW), found growing up to 15 cm in length (accession number PCK 041 -A, spicifera). This species has been reported from the Gulf of Mexico (Schneider & Searles 1991; Littler & Littler 2000), but has never been reported from the Texas Coastal Bend. This species is noted to be an early colonizer in areas of calm waters (Littler & Littler 2000), which is characteristic of the protected segment of the newly-dredged Packery Channel. This study proposes a range extension for the species to include the Upper Laguna Madre near Packery Channel. Hurricane Emily, a category five storm, opened Packery Channel prematurely on 22 July 2005, before completion of the jetties. Large-scale disturbance such as hurricanes are thought to deliver terrestrial-derived nutrients in concentrations sufficient to trigger and sustain macroalgal blooms (Cooper 1966; McCook 1999). Hurricanes often carry with them foreign species, trapped within eddies created by the storm. Increased storm activity over the past several years may further explain the initial presence of some tropical flora uncommon to the area. Literature Cited Britton, J. C. and B. Morton. 1989. Shore Ecology of the Gulf of Mexico. The University of Texas Press. 387 pp. 224 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Cheney, D. P. 1977. A new and improved ratio for comparing seaweed floras. J. Phycol. 13 (SuppL): 12 (Abstr.). Connell, J. H. & R. O. Slaty er. 1977. Mechanisms of succession in natural communities and their role in community stability and organization. Am, Nat. Ill: 1119-1144. Cooper, M. J. 1966. Destruction of marine flora and fauna in Fiji caused by the Hurricane of February 1965. Pacific Science 20:137-141. Edwards, P. & Kapraun, D.F. 1973. Benthic marine algae ecology in the Port Aransas, Texas area. Contrib. Mar. Sci. 17:15-52. Kapraun, D. F. 1980. Summer aspect of algal zonation on a Texas jetty in relation to wave exposure. Contrib. Mar. Sci. 23:101-109. Lehman, R. L, 1999. A checklist of benthic marine macroalgae from the Corpus Christ! Bay area. Texas!. Sci. 51:241-252. Littler, D. S. & M. M. Littler. 2000. Caribbean Reef Plants. Offshore Graphics, Inc. Washington, D.C. 542 pp. McCook, L. J. 1999. Macroalgae, nutrients and phase shifts on coral reefs: scientific issues and management consequences for the Great Barrier Reef Coral Reefs 18:357-367. Medlin, L. K. 1984. Short note on changes in the abundance and occurrence of six macroalgal species along the Texas coast of the Gulf of Mexico. Contrib. Mar. Sci. 27:85-91. Pedrini, A. 1984. Algas marinhas bentonicas do litoral Norte-Fluminense. I- Lista preliminar das algas “Arribadas” da Praia de Manguinhos (Guaxindiba - Sao Juao da Barra - Rio de Janeiro). Atas da Sociedade Botanica do Brasil, seccao Rio de Janeiro 2: 133-142. Schneider, C. W. and R. B. Searles. 1973. North Carolina marine algae. IT New records and observations of the benthic offshore flora, Phycologia 12: 201- 211. Schneider, C. W. and R. B. Searles. 1991. Seaweeds of the Southeastern United States: Cape Hatteras to Cape Canaveral. Duke University Press, Durham, 553 pp. Wardle, W. J. 1992. Range extension for Porphyra leucosticta Thuret (Rhodophyta: Bangiales) from the central to the upper Texas coast. Texas J. Sci. 44: 117-118. Wynne, M. J. 2005. A checklist of benthic marine algae of the tropical and subtropical western Atlantic: second revision. Gebruder Bomtraeger, Berlin, Germany, 152 pp. RLF at: E-mail: ryan.fikes@tamucc.edu TEXAS J. SCL 60(3), AUGUST, 2008 225 FIRST RECORD OF THE GRAY FOX (UROCYON CINEREOARGENTEUS) ON TEXAS BARRIER ISLANDS Gerrad D* Jones and Jennifer K, Frey Department of Fishery and Wildlife Sciences New Mexico State University, P,0. Box 30003, Campus Box 4901, Las Cruces, New Mexico 88003-8003 Barrier islands are dominant features of many coastal areas (Pilkey 2003). Approximately two-thirds of the Texas coastline is bordered by six major islands, the largest of these being Padre Island. Padre Island extends ca. 180 km from Corpus Christ! south to Port Isabel, and although it was once continuous, it is now divided into North and South Padre Islands by the Mansfield Channel. Although parts of Padre Island are extensively developed, Padre Island National Seashore is the largest (ca. 110 km) stretch of undeveloped barrier island in the world (Pilkey 2003). When compared to other island systems, relatively little research regarding mammals has been conducted on barrier islands. However, this research has demonstrated that barrier islands have relatively depauperate faunas when compared to adjacent mainland areas (McAlister & McAlister 1993; Hice & Schmidly 2002). In Texas, several species of mammals that exist in coastal mainland areas, including the gray fox {Urocyon cinereoargenteus), have not been reported from adjacent barrier islands (Hice & Schmidly 2002). While gray fox are widely distributed and are common throughout their range in Texas (Schmidly 2004), there have been no previous verified records of this species on Padre Island or other islands within the Gulf of Mexico (Selander et al. 1962; Hall & Dalquest 1963; Hice & Schmidly 2002). Bent (1940) reported observing gray fox on Cumberland Island, Georgia, but Ruckdeschel et al. (1986) later dismissed these accounts. This study reports the first confirmed record of the gray fox from Texas barrier islands. On 15 March 2005, National Park Service personnel found a dead female gray fox on North Padre Island, 1 1.0 km south of the end of Park Road 22 (Kleberg County, Padre Island National Seashore, Pan Am Road, 14 0664484E, 3023097N). The specimen was prepared as a 226 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Standard skin and skeleton voucher specimen (Padre Island National Seashore catalog number 3744). Measurements of this specimen were: total length, 960 mm; tail length, 402 mm; right hind foot, 129 mm; mass, 3.0 kg. Mammae showed no obvious signs of lactation. Seacoast bluestem {Schizachyrium scoparium) and gulf dune paspalum (Paspalum monostachyum) dominated the vegetation where the specimen was found. Since August 2005, five observations of gray fox have been made on Texas barrier islands (Jones 2008). On North Padre Island: 14 August 2005 (five specimens together), 14 August 2006 (one specimen), 9 September 2006 (one specimen). On South Padre Island: 15 August 2005 (one specimen), 17 may 2006 (one specimen). Gray fox also appear to be well established in residential areas at the north end of North Padre Island. Residents of Corpus Christi have regularly reported foxes in yards and on porches (Ellis pers. comm.). Crooks (2002) found that the relative abundance of gray fox in California was highest at the edge of urban areas. Gray fox are known to utilize and perhaps benefit from residential development through the use of anthropogenic resources (Harrison 1993; 1997) and it is possible that gray fox are taking advantage of available resources while avoiding predators in residential areas. Although the abundance of gray fox around residential areas on Padre Island is unknown, these individuals may be dispersing into surrounding undeveloped areas. Whatever the reason for their presence on Padre Island, it is possible that gray fox inhabit other barrier islands and have not yet been discovered. Acknowledgements We thank the staff at Padre Island National Seashore for all their help and logistical support, especially A. Wimer for salvaging the gray fox specimen and R. Larson and D. Echols for helpful information. We thank D. Echols and C. Ogden for reviewing a previous version of this paper. We thank Chad Stinson of the Aransas National Wildlife Refuge complex for helpful information about the distribution of gray fox in coastal Texas. We thank Mike Ellis of the Padre Island Moon for information on gray fox in Corpus Christi. We thank the National Park Service for funding this study (Cooperative Agreement Number CA- 1248-00-002). TEXAS J. SCI. 60(3), AUGUST, 2008 111 Literature Cited Bent, A. C. 1940. Life histories of North American cuckoos, goatsuckers, hummingbirds and their allies. U.S. Nat. Mus, Bull., 176:1-506. Crooks, K. R. 2002. Relative sensitivities of mammalian carnivores to habitat fragmentation. Conserv. Biol., 16:488-502. Hall, E. R. & W. W. Dalquest. 1963. The mammals of Veracruz. Univ. Kansas, Mus. Nat Hist, 14:165- 362. Harrison, R. L. 1993. A survey of anthropogenic ecological factors potentially affecting gray foxes {Urocyon cinereoargentens) in rural a residential area. Southwest. Nat., 38(4):352-356. Harrison, R. L. 1997. A comparison of gray fox ecology between residential and undeveloped rural landscapes. J. Wildl. Manage., 61:1 12-122. Hice, C. L. & D. J. Schmidly. 2002. The mammals of coastal Texas: A comparison between mainland and barrier island faunas. Southwest, Nat., 47(2):244-256. Jones, G. D. 2008. Mammalian zoogeography of Padre Island National Seashore, Texas. Unpublished M.S. thesis. New Mexico State Univ., Las Cruces, 266 pp. McAlister, W. H. & M. K. McAlister. 1993. Matagorda Island: a naturalist’s guide. Univ. Texas Press, Austin, 380 pp. Pilkey, F. 2003. A celebration of the world’s barrier islands. Columbia Univ. Press, New York, 400 pp. Ruckdeschel, C. & C. R. Shoop. 1986. The fauna of Cumberland Island I: on the occurrence of the opossum and gray fox. Georgia J. Sci., 44:90-95. Schmidly, D. J. 2004. The mammals of Texas: revised edition. Austin: Univ. Texas Press, 544 pp. Selander, R. K., R. F. Johnston, B. J. Wilks & G. G. Raun. 1962. Vertebrates from the barrier islands of Tamaulipas, Mexico. Univ. Kansas Publ., Mus. Nat. Hist., 12:309- 345. JKF at: jfrey@nmsu.edu FIRST REPORT OF THE BROWN ALGA PADINA GLABRA (OCHROPHYTA: DICTYOTALES) FROM THE COAST OF TEXAS AND THE GULF OF MEXICO Michael J. Wynne Department of Ecology and Evolutionary Biology and Herbarium University of Michigan, 3600 Varsity Drive, Ann Arbor, Michigan 48109 Up to now the only species of the brown algal genus Padina that has been recognized as occurring on the Texas coast has been P, gymnospora (Kiitz.) Sonder (Taylor 1960; Humm & Flilldebrand 228 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 1962; Earle 1969; Edwards 1970; Sorensen 1979, all as P. vickersiae). This species had been known as P. vickersiae Hoyt in Howe (Howe 1920), until Allender & Kraft (1983) recognized that the "'Padina gymnospord' of various authors had been mis¬ interpreted to be usually three cells in thickness, whereas the type {Zonaria gymnospora Kiitz.) is four cells thick in mid frond and 6-8 cells thick closer to the base, in agreement with Kutzing's (1859) original depiction. Therefore, Padina vickersiae Hoyt is a later taxonomic synonym of P. gymnospora. Some new species of red, green, and brown algae have been described from the Texas coast in the past few decades (Wynne & Edwards 1970; Wynne 1993; Scott et al. 2006) or newly reported species added to the marine algal flora (Baca et al. 1977; Kaldy 1977; De Yoe & Hockaday 2001; Strenth 2001; Kowalski et al. 2007). The primary source of new records has been from work carried out at the Flower Garden Coral Banks (National Marine Sanctuary) south of Galveston (Eiseman & Blair 1982; Gavio & Fredericq 2005). In the present study it was a matter of a more careful examination of Padina specimens from Texas in the University of Michigan Herbarium (MICH) that allowed for the recognition that two collections (one from Nueces County in south central Texas and one from Cameron County in southernmost Texas) were not the customary ''Padina gymnospord\ the common species on the Texas jetties, but a distinct species representing not only its first report from the coast of Texas but also from the Gulf of Mexico. Padina glabra Gaillard 1966: 226 Type locality -VoiniQ de Fann, Dakar, Senegal, West Africa. Holotype in the Museum National d'Histoire Naturelle, Paris (PC). Texas ~Vori Aransas jetty, Mustang Island, Nueces County, 27 July 1969, coll M. Wynne 2588 [MICH] (Fig. 1). Isla Blanca Beach State Park, jetty, South Padre Island, near Port Isabel, Cameron County, 24 March 1975, coll M. Wynne 4293 [MICH]. TEXAS J. SCL 60(3), AUGUST, 2008 229 Fig, 1. Padina glabra. Specimen from Port Aransas jetty, Texas (MICH 2588), Scale bar: 4 cm. Padina glabra was described from a site near Dakar, Senegal, and was characterized as the only species in the genus that lacked multicellular hairs that are typically arranged in concentric rows in other species of the genus (Gaillard 1966). The species was described as having nomlacerated blades attached by rhizoids, only 1.0" 1.5 cm tall, 3 or 4 cell layers in thickness, rarely with calcification, and with non-indusiate tetrasporangial sori arranged in concentric zones. The species was subsequently reported from Mandapam, southern India, by Rengasamy & Anand (1986), who reported blades to be 4. 0-5.0 cm tall, divided down the middle and bearing plantules. The important point is that their specimens 230 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 lacked hairs. Wynne & De Clerck (1999) made the first report for the occurrence of this species from the western Atlantic, namely, several collections from the vicinity of St. Augustine, St. Johns Country, Florida, U.S.A. Wynne & De Clerck (1999) also pointed out that Dangeard (1952) had reported on several collections of a hairless Padina (that he called "'Padina sp.”) from the very same location where Gaillard (1966) was later to describe P. glabra, Gaillard, however, did not cite the Dangeard (1952) publication. The Isla Blanca Beach State Park specimens, of which there were six, reached 8- 12 cm in height, and the Port Aransas species, of which there were two, reached up to 14 cm in height. The plants from Isla Blanca are much branched axes ending distally in rounded blades. The plants from Port Aransas seem more mature and are divided into elongate, lacerate segments (Fig. 1). Blades in both collections are usually four cell layers in thickness. Some sections show local regions of three cell layers in thickness. Calcification is either lacking or just barely present on the blades. The base of the plant is a well developed stupose portion reaching to 1 cm in thickness and with a spreading attachment area. Sporangial and oogonial plants were observed. In both collections, however, plants bear propagula. These are also referred to as plantules and brood buds (Thivy 1945). Such propagula have been reported in this same species by Rengasamy & Anand (1986) in Indian material and by Wynne & De Clerck (1999) in Florida material. They appear to develop from sporangia at a very young stage or possibly in place of sporangia. Such propagula have been noted in other species of the genus: in P. pavonica (Linn.) Thivy by Bitter (1899), P, durvillaei Bory de St.-Vincent and P. distromtica Hauck by Thivy (1945), P. gymnospora (Kutz.) Sonder by Hoyt (1920) and P. antillarum (Ktitz.) Piccone by Hauck (1887), Gaillard (1967), and Lawson & John (1977). Finally, in the Port Aransas plants the sporangia and propagula were observed to occur in two closely positioned rows on the inferior blade surface and in more irregularly arranged sori on the superior side of the blade. This pattern is essentially the same as the pattern present in P. TEXAS J. SCL 60(3), AUGUST, 2008 231 antillarum, in which the inferior surface of the blade bears sporangia in two rows straddling each line of hairs (Jaasund 1976, as P. tetrastromatica; Tseng 1983, as P. tetrastromatica; Wynne 1998; Wynne & De Clerck 1999). In the Port Aransas plants identified as P. glabra, however, these parallel rows of sporangia/propagula do not have hairs present separating them, A similar arrangement of propagula lying in two closely aligned rows present in Florida material of P. glabra was depicted by Wynne & De Clerck (1999: fig. 12). Literature Cited Allender, B. M. & G. T. Kraft. 1983. The marine algae of Lord Howe Island (New South Wales): the Dictyotales and Cutleriales (Phaeophyta). Brunonia, 6:73- 130. Baca, B. J., E. R. Cox & L. O. Sorensen. 1977. Observations on several benthic marine algae from South Padre Island, Texas. Southwest. Nat., 21 :459-462. Bitter, G. 1889. Zur Anatomic und Physiologic von Padina Pavonia. Ber. Deutsch. Bot. Gesellsch., 17:255-274, pL 20. Dangeard, P. 1952. Algues de la presqu’ile du Cap-Vert (Dakar) et de ses environs. Botaniste, 36:195-329. De Yoe, H. R. & D. L. Hockaday. 2001. Range extensions of Codium taylorii and Caulerpa proUfera from the Lower Laguna Madre, Texas. Tex. J. Sci., 53:190- 192. Earle, S. A. 1969. Phaeophyta of the eastern Gulf of Mexico. Phycologia, 7:71-254. Edwards, P. 1970, Illustrated guide to the seaweeds and sea grasses in the vicinity of Port Aransas, Texas. Contrib. Mar. Sci., 15 (Suppl.):l-128. Eiseman, N. J. & S. M. Blair. 1982. New records and range extensions of deepwater algae from East Flower Garden Bank, northwestern Gulf of Mexico. Contrib. Mar. Sci., 25:21-26. Gaillard, P. 1966. Un Padina nouveau des cotes d'Afrique: Padina glabra sp. nova. Phycologia, 5:222-226. Gaillard, P. 1967. Etude monographique de Padina tetrastromatica (Hauck). Bull. Inst. Fondamental Afr. Noire, ser. A, 29:447-463. Gavio, B. & S. Fredericq. 2005. New species and new records of offshore members of the Rhodymeniales (Rhodophyta) in the northern Gulf of Mexico. Gulf Mexico Sci., 2005:58-83. Hauck, F. 1887. Ueber einige von J. M. Hildebrandt in Rothen Meere und Indischen Ocean gesammelte Algen. Hedwigia, 26:41-45. Howe, M. A. 1920. Algae. Pp. 553-618, in The Bahama Flora (N. L. Britton & C. F. Millspaugh, published by the authors, New York, Hoyt, W. D. 1920. The marine algae of Beaufort, N. C., and adjacent regions. Bull. U.S. Bureau Fish. 36: 367-556, pis. 84-119. 232 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Humm, H. J. & H. H. Hildebrand. 1962. Marine algae from the Gulf coast of Texas and Mexico. Publ. Inst. Mar. Sci. [Port Aransas, Texas], 8:227-268. Jaasund, E. 1976. Intertidal Seaweeds in Tanzania. A Field Guide. University of Tromso, [Tromso, Norway], [iii] + 160 pp. Kaldy, J. E. 1997. Range extension of Halimeda incrassata (Chlorophyta, Bryopsidales): occurrence in the Lower Laguna Madre of Texas. Southwest. Nat., 41:419-423. Kowalksi, J. L., D. L. Hockaday, G, H. Boza, Jr. & H. R. De Yoe. 2007. Re¬ occurrence of the tropical green macroalga, Penicillus capitatus Lamarck (Chlorophyta: Bryopsidales), in Lower Laguna Madre, Texas. Tex. J. Sci., 59: 305-310. Kiitzing, F. T. 1859. Tabulae Phycologicae. Vol. 9. Nordhausen, viii + 42 pp., 100 pis. Lawson, G. W. & D. M. John. 1977. The marine flora of the Cap Blanc Peninsula: its distribution and affinities. Bot. J. Linn. Soc., 75:99-1 18. Rengasamy, R. & N. Anand. 1986. Studies on Padina glabra Gaillard. Phykos, 25:1-5. Scott, J. L., B. Baca, F. D. Ott & J. A. West. 2006. Light and electron microscopic observations on Erythrolobus coxiae gen. et sp. nov. (Porphyrideophyceae, Rhodophyta) from Texas U.S.A. Algae, 21:407-416. Sorensen, L. O. 1979. A Guide to the Seaweeds of South Padre Island, Texas. Illustrations by E. E. Pena. Gorsuch Scarisbrick Publ., Dubuque, Iowa, 123 pp. Strenth, N. E. 2001. Caiderpa proUfera (Chlorophyta: Caulerpaceae) from the Laguna Madre of South Texas. Tex. J. Sci., 53:187-189. Taylor, W. R. 1960. Marine Algae of the Eastern Tropical and Subtropical Coasts of the Americas. University of Michigan Press, Ann Arbor, Michigan, xi + 870 pp. Thivy, F. 1945. A revision of the genus Padina Adans. Unpublished Ph. D. dissertation, Univ. of Mich., Ann Arbor, 247 pp. Tseng, C. K. 1983. Common Seaweeds of China. Science Press, Beiing, x + 3 16 pp. Wynne, M. J. 1998. A study of Padina antillarum (Kiitzing) Piccone and a comparison with P. tetrastromatica Hauck (Dictyotales, Phaeophyta). Cryptogam., Algol., 4:271-289. Wynne, M. J, 1993. Prionitis pterocladina sp. nov. (Halymeniaceae, Rhodophyta), a newly recognized alga in the western Gulf of Mexico. Bot. Mar., 36:497-502. Wynne, M. J. & O. De Clerck. 1999. First reports of Padina antillarum and P. glabra (Phaeophyta, Dictyotaceae) from Florida, with a key to the western Atlantic species of the genus. Caribb. J. Sci., 35:286-295. Wynne, M. J. & P. Edwards. 1970. Polysiphonia boldii sp. nov. from Texas. Phycologia, 9:1 1-16. MJW at: mwynne@umich.edu TEXAS J. SCI. 60(3), AUGUST, 2008 233 REPRODUCTION IN THE SIAMESE LEAF-TOED GECKO, DIXONIUS SIAMENSIS {SQUAMATA: GEKKONIDAE) FROM THAILAND Stephen R, Goldberg Department of Biology, Whittier College, PO Box 634 Whittier, California 90608 The Siamese leaf-toed gecko (Siam-blattfmger), Dixonius siamensis is known from Thailand where it occurs from Songkhla, south to the Isthmus of Kra and north to at least Chiang Mai, from southern Myanmar to the Lao People’s Democratic Republic, Vietnam and likely Cambodia (Bauer et al. 2004). It occurs from sea level to at least 700 m (Manthey & Grossman 1997) and inhabits a variety of forest types and occasionally occurs in developed areas; females deposit clutches of two eggs during the rainy season (Cox et al. 1999). The purpose of this note is to provide information on the reproductive cycle of D. siamensis from a histological analysis of monthly gonad samples from museum specimens collected in Thailand. Minimum sizes for reproduction in males and females of S. siamensis are given. This paper represents the first detailed account of reproduction in D. siamensis. Comparisons are made with the timing of reproduction of other lizards from the tropics. A total of 105 D. siamensis from Thailand including 49 females (mean snout vent length, SVL = 42.5 mm ± 4.9 SD, range = 33-52 mm), 50 males (mean SVL = 41.5 mm ± 5.3 SD, range = 29-52 mm) and 6 presumed neonates (mean SVL = 21.0 mm ±1.1 SD, range = 19-22 mm) were examined from the herpetology collection of the Field Museum of Natural History (FMNH), Chicago, Illinois. Geckos were collected 1957-1960, 1967 and 1969. For histological examination, the left testis was removed from males and the left ovary was removed from females. Enlarged follicles (> 4 mm length) or oviductal eggs were counted. Tissues were embedded in paraffin and cut into sections of 5 pm. Slides 234 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 were stained with Harris hematoxylin followed by eosin counterstain (Presnell & Schreibman 1997). Slides of testes were examined to determine the stage of the spermatogenic cycle. Slides of ovaries were examined for the presence of yolk deposition or corpora lutea. Histology slides were deposited in the Field Museum of Natural History (FMNH) herpetology collection. An unpaired ^ test was used to compare D. siamensis male and female mean body sizes (SVL) using Instat (vers. 3.0b, Graphpad Software, San Diego, CA). The following specimens of D. siamensis from Thailand (by province) from the herpetology collection of the Field Museum of Natural History (FMNH), Chicago, Illinois, comprise the basis for this study: CHIANG MAI: 177752. CHON BURI: 177696, 177697, 177707, 177708, 177710, 177711, 177713, 177714, 177716, 177719, 177720, 177722-177725, 177728, 177731-177733, 177735, 177736, 177738, 177739-177741, 177743-177745, 181568, 181569, 181583, 181597. KANCHANABURI: 177698-177701, 177704. LOEI: 177747, 177748, 177762, 177763. NAKHON RATCHASIMA: 181474-181478, 181480-181484, 181486-181488, 181491, 181493-181495, 181497, 181498, 181537, 181538, 181543, 181547, 181553, 181555, 181561, 181562, 181571, 181573-181576, 181580, 181585, 181589, 181593, 181596, 181602-181604, 181606, 181608, 181609, 181614-181616, 181625, 181627, 181630, 181633-181637, 181640, 181641, 181644, 181645, 181647, 181652, 181664. PRACHUAP KHIRIKHAN: 177694. UBON RATCHATHANI: 177695. There was no significant size difference (mean SVL) between Z). siamensis males and females (unpaired ^test). Two stages in the testicular cycle were observed. Recrudescence in which cellular renewal for the next period of sperm formation (= spermiogenesis) occurs, was typified by spermatogonia and primary spermatocytes. Spermiogenesis (sperm production) was characterized by groups of spermatozoa and metamorphosing spermatids lining the lumina of the seminiferous tubules. The presence of males undergoing spermiogenesis in twelve months of the year (Table 1) indicates a continuous testicular cycle. Males in recrudescence were found TEXAS J. SCI. 60(3), AUGUST, 2008 235 Table 1. Monthly stages in testicular cycle of Dixonius siamensis from Thailand. Month n Recrudescence Spermiogenesis January 5 0 5 February 2 0 2 March 4 0 4 April 5 0 5 May 3 0 3 June 2 0 2 July 6 1 5 August 2 1 1 September 5 0 5 October 9 0 9 November 3 0 3 December 4 0 4 only in July (FMNH 177701, SVL = 33 mm) and August (FMNH 181537, SVL = 36 mm). However, samples of D. siamensis from these months are too small to speculate on the significance of this finding, i.e., whether they might suggest a “resting” period in the testicular cycle. The smallest reproductively active male (FMNH 181641) measured 29 mm SVL and was collected in April. Mean clutch size for 14 gravid females was 1.86 ± 0.36 SD, range = 1-2. Females with either enlarged ovarian follicles (> 4 mm), soon to be ovulated, or oviductal eggs were found in all months except September to December (Table 2). Nevertheless, the presence of females exhibiting early yolk deposition (basophilic yolk granules within the follicle) in September, October and November suggests ovarian activity is still occurring, but perhaps at a slower rate than January to August. The smallest reproductively active female (FMNH 181627), collected in June, measured 37 mm SVL and was undergoing early yolk deposition. One female from April (FMNH 181495) that exhibited no yolk deposition contained a 236 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Table 2. Monthly stages in the ovarian cycle of Dixonius siamensis from Thailand. Month n No yolk Deposition Early yolk deposition Follicles > 4 mm Oviductal eggs January 3 1 1 0 1 February 3 2 0 1 0 March 6 1 2 0 3 April 6 2* 1 1 2 May 2 1 0 0 1 June 4 0 2 0 2 July 3 0 1 1 1 August 3 1 0 1 1 September 4 2 2 0 0 October 10 9 1 0 0 November 3 3 0 0 0 December 2 1 1 0 0 * one female contained a corpus luteum. corpus luteum indicating an egg clutch had recently been deposited. There was no evidence (corpus luteum and/or oviductal egg) and concomitant yolk deposition to indicate that D. siamensis produces more than one clutch in the same year. However, this may reflect small sample sizes. Presumed neonates were collected in July {n = 1, SVL 20 mm ), September {n = 4, SVL = 20, 21, 21, 22 mm) and October {n=\, SVL =19 mm). The reproductive cycle of D. siamensis appears similar to that of other tropical lizards as it exhibits an extended period of sperm formation and egg production (Fitch 1982). This pattern appears typical for other gekkonid species studied in the tropics as reported for Cosymbotus platyurus (currently Hemidactylus platyurus), Hemidactylus frenatus and Peropus mutilatus (currently Gehyra mutilata) from West Java, Indonesia (Church 1962) and Cyrtodactylus malayanus and C. pubisulcus from Borneo (Inger & Greenberg 1966). In contrast, gekkonids from temperate areas TEXAS J. SCI. 60(3), AUGUST, 2008 237 exhibit a seasonal cycle with most reproduction occurring in spring (Flemming & Bates 1995; Goldberg 1997; Goldberg 2006a; Goldberg 2006b). This pattern is seen in many other species of temperate zone lizards (see for example, Goldberg 1974; Goldberg 1975) indicating the local environment plays a role in adjusting life history characteristics of lizards (Vitt 1986). Dixonius siamensis also produces small clutches of eggs (1-2) as is typical for other gekkonid species (Vitt 1986). Kluge (1967) categorized gecko reproductive cycles as: (1) no definite seasonal reproductive cycle and mating may occur throughout the year, and (2) breeding is cyclic and restricted to a short period during the year. Dixonius siamensis clearly fits into the former category. However, with at least 39 species of geckos recorded from Thailand (Chan-Ard et al. 1999), subsequent studies are needed before the variations in their reproductive cycles can be ascertained. Acknowledgments I thank Maureen M. Kearney and Alan Resetar (Field Museum of Natural History), Chicago, Illinois for permission to examine D. siamensis. Literature Cited Bauer, A. M., M. Sumontha, W. Grossmann, O. S. G. Pauwels & G. Vogel. 2004. A new species of Dixonius (Squamata: Gekkonidae) from Kanchanaburi Province, Western Thailand. Current Herpetology, 23 : 1 7-26. Chan-Ard, T., W. Grossmann, A. Gumprecht & K. Schulz. 1999. Amphibians and reptiles of peninsular Malaysia and Thailand. An illustrated checklist. Bushmaster Publications, Wuerselen, Germany, 240 pp. Church, G. 1962. The reproductive cycles of the Javanese house geckos, Cosymbotus platyurus, Hemidactylus frenatus, and Peropus mutilatus. Copeia, 1962:262-269. Cox, M. J., P. P. van Dijk, J. Nabhitabhata & K. Thirakhupt. 1999. A photographic guide to snakes and other reptiles of peninsular Malaysia, Singapore and Thailand. Ralph Curtis Publishing Co., Sanibel Island, Florida, 144 pp. Fitch, H. S. 1982. Reproductive cycles in tropical reptiles. Mus. Nat. Hist. Univ. Kansas, Occas. Pap., 96:1-53. 238 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 Flemming, A. F. & M. F. Bates. 1995. Male and female reproductive cycles of Bibron’s gecko Pachydactylus bibronii (Sauria: Gekkonidae) in the Free State province of South Africa. J. Afr. ZooL, 109:409-422. Goldberg, S. R. 1974. Reproduction in mountain and lowland populations of the lizard Seel oporus occidental is. Copeia, 1974:176-182. Goldberg, S. R. 1975. Reproduction in the sagebrush lizard, Sceloporus graciosus. Amer. Midi. Nat., 93:177-187. Goldberg, S. R. 1997. PhyUodactyliis xanti (Leaf-toed Gecko). Reproduction. Herpetol. Rev., 28:152-153, Goldberg, S. R. 2006a. Reproductive cycle of the Namib giant ground gecko, Chondrodactylus angidifer (Squamata: Gekkonidae). Afr. Zook, 41 :308-3 11. Goldberg, S. R. 2006b. Note on reproduction of Wahlberg’s Kalahari gecko, Col opus wahlbergii (Squamata: Gekkonidae) from southern Africa. Bull. Maryland Herpetol. Soc., 42:140-143, Inger, R. F. & B. Greenberg. 1966. Annual reproductive patterns of lizards from a Bornean rain forest. Ecology, 47 : 1 007- 1021. Kluge, A. G. 1967. Higher taxonomic categories of gekkonid lizards and their evolution. Bull. Amer. Mus. Nat. Hist., 135(1): 1-59. Manthey, U. & W. Grossmann. 1997. Amphibien & Reptilien Sudostasiens. Natur und Tier Verlag Munster, Germany, 512 pp. Presnell, J. K. & M. P. Schreibman. 1997. Humason’s Animal Tissue Techniques. The Johns Hopkins University Press, Baltimore, 572 pp. Vitt, L. J. 1986. Reproductive tactics of sympatric gekkonid lizards with a comment on the evolutionary and ecological consequences of invariant clutch size. Copeia, 1986:773-786. SRG at: sgoldberg@whittier.edu THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 239 IN RECOGNITION OF THEIR ADDITIONAL SUPPORT OF THE TEXAS ACADEMY OF SCIENCE DURING 2008 PATRON MEMBERS Stephen R. Goldberg Deborah D. Hettinger Don W. Killebrew David S. Marsh Ned E. Strenth SUSTAIND^G MEMBERS James Collins Donald E. Harper, Jr. Michael Looney Ernest L. Lundelius, Jr. John Sieben Fred Stevens William F. Thomann Milton W. Weller SUPPORTING MEMBERS Michael L. Morrison Jim Neal William J. Quinn Lynn Simpson Renard Thomas 240 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 3, 2008 THE TEXAS ACADEMY OF SCIENCE www.texasacademyofscience.org Membership Information and Application MEMBERSHIP.-Any person or member of any group engaged in scientific work or interested in the promotion of science is eligible for membership in The Texas Academy of Science. 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No part of this publication may be reproduced, stored in a retrieval system or transmitted, in any form or by any means, electronic, mechanical, recording or otherwise, without the prior permission of the Managing Editor of the Texas Journal of Science. THE TEXAS JOURNAL OF SCIENCE Volume 60, No. 4 November, 2008 CONTENTS Riparian Bird Community from the Rio Sabinas, Coahuila, Mexico. By Irene Ruvalcaba-Ortega, Jose /. Gonzdlez-Rojas and Ricardo Canales-del Castillo . . . . . . . .243 Reproduction in Seven Species of Microlophus (Squamata: Tropiduridae) from South America. By Stephen R. Goldberg . . . . . 261 Notes on Food Attraction and the Demographics of Blue Crab {Callinectes sapidus) Trapping in Tres Palacios Bay, Texas. By Joel D. Anderson and Dusty L. McDonald . . . 27 1 Spatial and Temporal Distributions of Planktonic Diatoms in a Subtropical Bayou, along the Upper Texas Coast. By Antonietta Quigg and Linda Roehrborn . 28 1 An Evaluation of Modified IMS Swabs for the Screening of Oxidizers and Home-Made Explosives. By Karl D. Kuklenz and Darren L. Williams . 299 General Notes Notes on Behavior of the Texas Kangaroo Rat {Dipodomys elator). By Jim R. Goetze, Allan D. Nelson and Chad Stasey . 309 Reproductive Cycle of the Brown Forest Skink, Sphenomorphus cherriei (Squamata: Scincidae), from Costa Rica. By Stephen R. Goldberg . 317 The Distribution of Spotted Skunks, Genus Spilogale, in Texas. By Robert C. Dowler, Carla E. Ebeling, Gema L Guerra and Adam W. Ferguson . . . 321 Index to Volume 60 (Subject, Authors & Reviewers)..... . 327 Recognition of Member Support . . . . . . . . . 333 Membership Application . 334 Postal Notice 335 THE TEXAS JOURNAL OF SCIENCE EDITORIAL STAFF Managing Editor: Ned E. Strenth, Angelo State University Manuscript Editor: Frederick B. Stangl, Jr., Midwestern State University Associate Editor for Botany: Janis K. Bush, The University of Texas at San Antonio Associate Editor for Chemistry: John R. Villarreal, The University of Texas-Pan American Associate Editor for Computer Science: Nelson Passos, Midwestern State University Associate Editor for Environmental Science: Thomas LaPoint, University of North Texas Associate Editor for Geology: Ernest L. Lundelius, University of Texas at Austin Associate Editor for Mathematics and Statistics: E. Donice McCune, Stephen F. Austin State University Associate Editor for Physics: Charles W. Myles, Texas Tech University Manuscripts intended for publication in the Journal should be submitted in TRIPLICATE to: Dr. Frederick B. Stangl, Jr. TJS Manuscript Editor Department of Biology Midwestern State University Wichita Falls, Texas 76308 frederick.stangl@mwsu.edu Scholarly papers reporting original research results in any field of science, technology or science education will be considered for publication in The Texas Journal of Science. Instructions to authors are published one or more times each year in the Journal on a space-available basis, and also are available on the Academy's homepage at: www.texasacademyofscience.org AFFILIATED ORGANIZATIONS American Association for the Advancement of Science, Texas Council of Elementary Science Texas Section, American Association of Physics Teachers Texas Section, Mathematical Association of America Texas Section, National Association of Geology Teachers Texas Society of Mammalogists TEXAS J. OF SCIENCE 60(4):243-260 NOVEMBER, 2008 RIPARIAN BIRD COMMUNITY FROM THE RIO SABINAS, COAHUILA, MEXICO Irene Ruvalcaba-Ortega, Jose I. Gonzalez-Rojas and Ricardo Canales-del Castillo Laboratorio de Biologia de la Conservacion, Facultad de Ciencias Biologicas Universidad Autonoma de Nuevo Leon, Apartado Postal 25-F Cd. Universitaria, San Nicolas de los Garza Nuevo Leon, Mexico 66451 Abstract -Riparian areas have been identified as key habitats for wildlife, especially for the North American arid landscape; however, similar areas in northern Mexico have been poorly studied. This study was conducted in a riparian habitat dominated by Montezuma bald cypresses (Taxodium mucronatum) of the Rio Sabinas, Coahuila, Mexico, from December 2004 to November 2005. Monthly samples were conducted, using a total of 443 point counts, in three river sections characterized by distinct levels of disturbance, from lesser to greater: Melchor Miizquiz, San Juan de Sabinas and Sabinas. A total of 168 species were recorded. The Melchor Muzquiz section exhibited the smallest number of species corrected by rarefaction and a significantly smaller number of species and individuals per point count. The Olmstead-Tukey diagrams per section also showed differences in the dominance of the species. From a conservation point of view, noteworthy species were recorded such as Wood Stork and Painted Bunting. Resumen.-Las zonas riberenas han sido identificadas como habitats clave para la vida silvestre, especialmente para el paisaje arido de Norteamerica; sin embargo en el norte de Mexico dichas zonas han sido poco estudiadas. El presente estudio se llevo a cabo en el habitat ribereno dominado por sabinos {Taxodium mucronatum) del Rio Sabinas, Coahuila, Mexico, de diciembre de 2004 a noviembre de 2005. Se llevaron a cabo muestreos mensuales utilizando un total de 443 puntos de conteo de radio fijo en tres zonas del rio caracterizadas por distintos niveles de disturbio, de menor a mayor: Melchor Muzquiz, San Juan de Sabinas y Sabinas. Se registraron un total de 168 especies. La zona Melchor Muzquiz mostro el menor numero de especies corregidas por rarefaccion y un numero significativamente menor de especies e individuos por punto de conteo. Los diagramas Olmstead-Tukey por zona tambien mostraron diferencias en la dominancia de las especies. Se registraron especies importantes desde el punto de vista de la conservacion, como Mycteria americana y Passerina ciris. The importance of riparian zones for breeding and migrant birds has been extensively documented (e. g., Knopf et al. 1988; Knopf & Samson 1994; Skagen et al. 1998; Anthony et al. 1996; Finch & Yong 2000; Flannery et al. 2004; Kelly & Hutto 2005; Villasenor 244 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 2006), While riparian woodlands of North America constitute less than 1% of the area of arid landscapes they support more reproductive birds than the extensive surroundings (e.g., Knopf et al. 1988). Unfortunately, it has been suggested that at least 95% of all western riparian habitats have been altered in some way during the past century (Ohmart 1994), and that such destruction may be the most important factor in the decline of some western North American landbird species during the past century (DeSante & George 1994). However, the effects of such alterations on riparian systems may be unnoticed for decades due to longevity of cottonwoods (Rood & Mahoney 1993) and other riparian trees such as Montezuma bald cypresses. This could cause a delay in awareness of the need for conservation, management, and restoration actions. In northwestern Mexico, there have been few studies conducted on the importance of riparian habitats for migratory birds (Hutto 1995; Villasenor 2006), but none in the northeastern portion. The Rio Sabinas (130 km length) is located in northeastern Coahuila, running from the Sierra Santa Rosa near Melchor Muzquiz to the Venustiano Carranza Dam near Juarez municipality (Fig. 1). Its avifauna has been little studied even though it is a Priority Terrestrial Region for Conservation in Mexico (RTP-152, Arriaga et al. 2000), a State Natural Protected Area (Periodico Oficial 1998), and an important biological corridor linking northern Mexico with the southern United States through the well-conserved Sierra Santa Rosa, Maderas del Carmen and Big Bend National Park. This connection is also maintained downstream from the Venustiano Carranza Dam where the river is named the Rio Salado; it flows through Nuevo Leon and Tamaulipas and enters the Falcon Reservoir which is part of the Rio Grande. Its riparian vegetation is dominated by Montezuma bald cypress (Taxodium mucronatum, locally known as “sabino”), while pecan (Carya illinoinensis), sycamore (Platanus occidentalis), sugar hackberry (Celtis laevigata), and Mexican ash (Fraxinus berlanderiana) are also present. Its shrub stratum is variable but includes spiny hackberry RUVALCABA-ORTEGA, & GONZALEZ-ROJAS & CANALES-DEL CASTILLO 245 lorso'o'w ioi"0’0"w loo^so'o'w Figure 1. Map of the Rio Sabinas, Coahuila, Mexico, (C pallida), huisache {Acacia farnesiana), catclaw {A. greggii), blackbrush {A. rigidula), honey mesquite {Prosopis glandulosa), Texas persimmon {Dyospiros texand), and Mexican mulberry {Morus celtidifolia). The herbaceous cover is composed principally of straggler daisy {Calyptocarpus vialis). This area faces serious conservation issues such as lack of biological knowledge, overgrazing, deforestation, and open-ceiling coal mining. The latter is especially harmful because it drastically modifies the environment and frequently breaks into the freatic mantle and extracts water (sometimes from very near the river: Trevifio-Garza et al. 2002), affecting the hydrologic dynamics of the basin and interrupting the water flow in some sections of the river. Three distinct sections are observed in the riparian habitat qualitatively characterized by different levels of disturbance. The Melchor Miizquiz (MM) municipality section has a well preserved riparian habitat with mature and young woodland patches; the San Juan de Sabinas (SJS) municipality section contains partially preserved habitat, with mature patches but several rural and urban 246 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 localities; and the Sabinas (S) municipality section has deteriorated and fragmented riparian woodland vegetation in a narrow corridor which includes dead bald cypresses (due to coal mining activities). Therefore, the objectives of this study were to characterize the bird community of the Rio Sabinas and compare communities among sections with different levels of disturbance. Methods A systematic sampling of birds was conducted from December 2004 to November 2005 along the Rio Sabinas, in localities where Montezuma bald cypresses were present near the municipalities of Melchor Miizquiz (MM), San Juan de Sabinas (SJS) and Sabinas (S), Sections were visited monthly on a rotating basis to obtain a balanced sampling design. A total of 443 point counts (10 min, 25 m radius: Ralph et al. 1996) in 247 randomly selected sites were sampled, covering an area of 34.5 ha and 64 km of river length (approximately 50% of the total). Point counts were conducted by a single observer to reduce bias in detectability. The area was visited from December 2005 to January 2006 and May to August 2006, adding new records to the check-list. Species, including common and scientific names, were listed in phylogenetic order (Tables 1 & 2) following the AOU checklist (1998; 2000) and its supplements (Banks et al. 2002-2007). The residency status of each species was determined on the basis of field observations and information provided by Howell & Webb (1995). Only results from the one-year systematic effort were used for the analysis described below. To demonstrate that the sampling effort generated species accumulation curves reached the asymptote, the Estimates program (Colwell 2006) was used (Mao TauN Model, Colwell et al. 2004). Log-normal (truncated) and log-series models were assessed for goodness-of-fit (Kolmogorov-Smimov test) to the observed species abundance distribution of the whole community (e.g., Fisher et al. 1943; Preston 1948). RUVALCABA-ORTEGA, & GONZALEZ-ROJAS & CANALES-DEL CASTILLO 247 Density (D) was calculated as the number of birds per hectare. Frequency of occurrence (FO) was also obtained based on the percentage of point counts at which each species was observed. In both variables, a total of 443 point counts was considered for permanent resident and occasional species, 234 point counts for winter residents and transients (Oct-Mar), and 209 point counts for summer residents (Apr- Aug). In order to rank the species by dominance, two Olmstead-Tukey Diagrams were used for the whole community (Sokal & Rohlf 1995), one for species present during the breeding season (Apr-Aug) and one for those observed during the migratory-wintering season (Sep-Mar). The average values of FO and abundance (expressed as In + 1) are used as discriminators for four classes. Common (or dominant) species are those with values equal to or greater than abundance and FO averages. Frequent (or constant) species are those with a value less than the abundance average, but another equal to or greater than FO average. Uncommon (or occasional) species are those with a value less than the FO average, but another equal to or greater than abundance average. Finally, rare species are those with values less than both averages. Olmstead-Tukey Diagrams were used to compare communities among sections during the breeding season. Also, for an unbiased comparison of number of species, rarefaction curves (±95%CI; Mao Tau’s Model, Colwell et al. 2004) were obtained for each section using Estimates (Colwell 2006). Finally, Shannon’s Diversity Index (1948) and the Bray-Curtis Index of Similarity (1957) were calculated for each section and for the entire community. Results And Discussion A total of 168 species were observed. Of these, 58 were con¬ sidered as permanent residents, 32 as summer residents, 40 as winter residents, 30 as transients and nine as occasional. Eight confirmed and three potentially new breeding, and nine new non¬ breeding species were recorded for Coahuila (further discussed in Ruvalcaba-Ortega & Gonzalez-Rojas 2009), emphasizing the little 248 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 6 I c/> C 2 -= S O C/3 t/3 « u a, 12 ^ c/5 1> ^ o. .lb c I I s o C'-l si 2 O t> O 4> « V. z Q ^ w m % . r. C o ^ o Q o C/3 P H k od J fc a> O, ^ !! OC Cl, -O c3 ^ i -o c 5> fc 11 o “ I II 2 ^ O' 1 & o tu o . ^ — ' ss II :s 3 2 ^ c/^ tl 11 ' H (/5 O ' w 00 c .a It' S S ' 05 w o-,2 0.^0 c h- a> v .2 £ -o C 3 3 0 JC H Q < H > O H m o ” CJ QsS oS ai Qs: . 00 S C- — '.et r*n rn «o rs <2 OS OS TT — r4 cN •vf rn f6 rsj 0^ :d ZD :z c ci- kr ao ^ z V5 '!3- O' <2 in:;) u X cc X X X X > VO ^ X X =) X ’ZD zt ZD tXXXXZDZDUX > ■'fovvo«r5r'«a©«'ooooe5*-io o X X X X XXXXXXXXXXXXXX cua.crfi.a.(/5& P ^ ©«.« O «/ O S a» S I. Q tc s_ 0/ O ^ U O ^ 2 ^ *2 2 Qg y t g § *s s. § g I .y t ;l ^ ^ ^ ^ 1 ^ ■S'® §■ I I I si- 15 1^ S M S S £ .s ? £ a. 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I s. al c« c« Y = fe J O J a 1 3 *5 CO 00 &, o 2 "H ^ U I "S • I « s: U « 5 X. -f ^ ^ I « o ® ^ U O CQ CO ii o o *JI o S 00 S M -o 3 3 Q < K I f .Sj S ^ J 5 I* S .a ■s « ? i I & •I m I i If fc*^S 5453 § § ©40 251 Species under Mexican law protection (NOM-059-SEMARNAT-2001, DOF 2002); Species listed as threatened for Texas (TPWD 2003); New records for Coahuila (Ruvalcaba-Oitega & Gonz^Iez-Rojas 2009). 252 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Table 2. List of non-breeding species in phylogenetic order (AOU 1998, 2000; Banks et al. 2002-2007); their residency status (R): WR=Winter Resident; T=Transient; and 0=Occasionai. Density (D, birds/hectare) is shown per species. Olmstead Tukey Dominance for wintering season (OT-W): C=Common; U= Uncommon; R=Rare. Species in bold were used for quantitative analysis. R D OT-W Scientific Name Common Name Non-breeding Species Abe sponsa Wood Duck WR/PR? 0.45 U Anas strepem Gadwall WR 0.14 R Anas discors Blue-winged Teal T 0.24 U Anas cyanoptera Cinnamon Teal T Anas crecca Green- winged Teal T Phalacrocorax auritus Double-crested Cormorant T Egretta tricolor Tricolored Heron O 0.02 R Nycticorax nycticorax Black-crowned Night-Heron WR 0.10 R Plegadis chihP White-faced Ibis T Mycteria americana*°^ Wood Stork 0 Circus cyaneus Northern Harrier O Accipiter striatus* Sharp-shinned Hawk WR 0.07 R Accipiter cooperii* Cooper’s Hawk WR 0.07 R Buteo albicaudatus*° White-tailed Hawk O 0.02 R Falco sparverius American Kestrel T 0.03 R Falco columbarius Merlin WR? Grus canadensis* Sandhill Crane T? 0.07 R Actitis macularius Spotted Sandpiper WR 0.95 C Calidris minutilla Least Sandpiper WR 0.17 u Gallinago delicata Wilson’s Snipe WR Zenaida asiatica White- winged Dove 0? Ara militarist Military Macaw O? Megaceryle alcyon Belted Kingfisher WR 0.54 c Sphyrapicus varius Yellow-bellied Sapsucker WR 0.17 u Sphyrapicus nuchalis Red-naped Sapsucker WR 0.07 R Colaptes auratus Northern Flicker WR 0.03 R Contopus cooperi Olive-sided Flycatcher T 0.14 R Contopus sordidulus Western Wood-Pewee T 0.03 R Contopus Virens Eastern Wood-Pewee T Empidonax sp. Flycatcher T 0.07 R Empidonax minimus Least Flycatcher T Empidonax wrightii Gray Flycatcher WR Empidonax oberholseri Dusky Flycatcher T Sayornis phoebe Eastern Phoebe WR 1.90 C Sayornis saya Say's Phoebe WR Tyrannus forficatus Scissor-tailed Flycatcher T Lanins ludovicianus Loggerhead Shrike WR? 0.03 R Vireo vicinior Gray Vireo T? 0.07 R Vireo cassinii Cassin’s Vireo T 0.03 R Vireo solitarius Blue-headed Vireo WR 0.14 R RUVALCABA-ORTEGA, & GONZALEZ-ROJAS & CANALES-DEL CASTILLO 253 Table 2 cont. Scientific Name Common Name R D OT-W Non-breeding Species Stelgidopteryx serripennis Northern Rough-winged SwaUow WR/PR? 0.61 C Troglodytes aedon House Wren WR 0.78 c Cistothorus palustris Marsh Wren WR Regulus satrapa Golden-crowned Kinglet WR 0.24 u Regulus calendula Ruby-crowned Kinglet WR 2.58 c Sialia sialis Eastern Bluebird WR 0.58 u Catharus guttatus Hermit Thrush WR 0.03 R Turdus grayf Clay-colored Robin O? Turdus migratorius American Robin WR Anthus rubescens American Pipit WR Bombycilla cedrorum Cedar Waxwing T 0.48 U Vermivora celata Orange-crowned Warbler WR 0.41 U Vermivora ruficapilla Nashville Warbler T 0.14 R Dendroica petechia Yellow Warbler T 0.10 R Dendroica coronata Yellow-rumped Warbler WR 5.27 C Dendroica nigrescens Black-throated Gray Warbler T 0.03 R Dendroica virens Black-throated Green Warbler T 0.03 R Dendroica townsendi Townsend’s Warbler T 0.07 R Dendroica occidentalis Hermit Warbler T Dendroica dominica Yellow-throated Warbler O 0.02 R Dendroica pin us Pine Warbler O 0.02 R Mniotilta varia Black-and-white Warbler WR 0.07 R Seiurus aurocapilla Ovenbird T 0.03 Seiurus noveboracensis Northern Waterthrush T Oporornis tolmiei* MacGillivray’s Warbler T 0.03 R Wilsonia pusilla Wilson’s Warbler WR 0.24 U Piranga ludoviciana Western Tanager T 0.14 R Spizella passerina Chipping Sparrow WR Spizella pallida Clay-colored Sparrow WR 0.03 R Spizella pusilla Field Sparrow WR 0.78 U Pooecetes gramineus Vesper Sparrow WR Passerculus sandwichensis Savannah Sparrow WR Melospiza melodia Song Sparrow WR? Melospiza lincolnii Lincoln’s Sparrow WR 1.29 c Melospiza georgiana Swamp Sparrow WR 0.37 u Zonotrichia leucophrys White-crowned Sparrow WR 0.24 u Pheucticus melanocephalus Black-headed Grosbeak T Icterus galbula Baltimore Oriole T Carduelis pinus Pine Siskin T 0.14 R * Species under Mexican law protection (NOM-059-SEMARNAT-2001, DOF 2002); ° Species listed as threatened for Texas (TPWD 2003); New records for Coahuila (Ruvalcaba-Ortega & Gonzalez-Rojas 2009). 254 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO, 4, 2008 attention previously given to this area. Fifteen of the species are under protection by Mexican laws (NOM-059-SEMARNAT-2001, DOF 2002), such as Muscovy Duck, Wood Stork, and Red¬ shouldered Hawk. In Texas as well, six of the species are listed as threatened for the state, e.g.. White-faced Ibis, Gray Hawk, and Northern Beardless-Tyrannulet (TPWD 2003). Painted Bunting, a near threatened species on the Red List of Endangered Species (lUCN 2007), was also recorded as a common summer breeder in the area (Table 1). Based only on the systematic sampling effort, 126 species (75% of the total) were recorded and used for ftirther quantitative analysis. The accumulation curve of observed and expected species reached the asymptote assuring a representative sampling (Fig, 2). The species abundance distribution for the whole community was best fitted by the log-series distribution (D=0.035, P>0.05), which is suggested to be associated with a community dominated by one or a few factors (Magurran 2004). This would be expected for a restricted vegetation type, such as the riparian woodlands of northern Mexico, which are generally immersed in a matrix of semi-arid scrubland. During the breeding season, 40 common, 12 uncommon, and 42 rare species were observed for the Rio Sabinas bird community (Table 1). The most dominant species were the Carolina Wren (D=5.28 birds/ha; A=173 inds.; FO= 62.2%) and Golden-fronted Woodpecker (D=5.19 birds/ha; A= 198 inds.; FO=57.9%) as permanent residents, and Summer Tanager (D=6.36 birds/ha; A=167 inds.; FO=50.2%), as a summer breeder. During the migrating/wintering season, 21 common, 21 uncommon, and 59 rare species were observed (Table 2). The most dominant species were permanent residents such as the Carolina Wren (A=93 inds.; FO= 48.2%), Black-crested Titmouse and Northern Cardinal (both D=0.18 birds/ha; A=76 inds.; FO=39,8%), but wintering residents were also present among common species, e. g,, Yellow-rumped Warbler (D=2.78 birds/ha; A=155 inds.; FO=26.5%) and Ruby- crowned Kinglet (D=1.37 birds/ha; A=76 inds.; FO=23.5%). RUVALCABA-ORTEGA, & GONZALEZ-ROJAS & CANALES-DEL CASTILLO 255 Figure 2. Observed (solid line) and expected (bold line) ±95%CI (dotted lines) species accumulation curves of the Rio Sabinas bird community sampled from December 2004 to November 2005. Important differences among communities were found when separating the analysis by sections. During the breeding season, MM (43 species, 694 individuals) showed the Carolina Wren (A=102 inds.; FO= 68%) and Summer Tanager (A=98 inds.; FO= 50%) as the dominant species; SJS (49 species, 800 individuals) also showed the Carolina Wren as the most dominant species (A=74 inds.; FO= 53.4%) but included the Golden- fronted Woodpecker (A=75 inds.; FO= 52.3%) as the second. On the other hand, S (58 species, 736 individuals) showed a community mostly dominated by the Golden-fronted Woodpecker (A=102 inds.; FO= 82.43%), a species that adjusts well to human-altered environments (Husak & Maxwell 1998). Also, the Painted Bunting was determined to be a common species only for the MM section (A=20 inds.; FO= 14%), where as SJS (A=7 inds.; FO= 8%) and S (A=7 inds.; FO= 8.1%) included this species as uncommon (almost rare). Rarefacted species richness for MM was significantly lower than SJS and S; however SJS and S were not distinct from one another (Fig. 3). A Kruskal- Wallis test among sections, comparing number of species and individuals per point count, showed highly 256 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Figure 3. Rarefacted (by sample) species accumulation curves (±95%CI) for each riparian section: MM (bold line), SIS (solid line), S (dashed line) in the Rio Sabinas, Coahuila (Mao Tau's Model, Colwell et al 2004). significant differences in both cases (richness: KW2,443 = 39.61, p< 0.01; abundance: KW2,443 = 27.90, p< 0.01). A posterior Mann- Whitney test between each section showed statistically significant differences in all cases, showing the highest mean number of species and individuals for S, and the smallest for MM (Richness: MM vs. SJS, U=10332, p<0.05; MM vs. S, U=5418, p<0.01; SJS vs. S, U=6095, p<0.01; abundance: MM vs, SJS, U==10196, p<0.05; MM vs. S, U=5789, p<0.01; SJS vs. S, U-6645, p<0.01). Higher numbers of species and individuals in S could be a consequence of a more open habitat with better visibility and a higher detection rate. Also, a more fragmented habitat with a greater influence of arid scrubland species such as the Scissor-tailed Flycatcher, Loggerhead Shrike, and Pyrrhuloxia, which were recorded only in this section. Using Bray-Curtis’ Index, the highest value of similarity was obtained for the S and SJS sections (66.3%), followed by MM with S (63.7%) and SJS (63.5%). Diversity values showed the same pattern across sections, the highest was S (H' = 3.8, J’=0.84), followed by SJS (H’ = 3.6, J’ =0.83), and M (H’ = 3.5, J’=0.82). The whole community showed a higher diversity (H'=3.8), but a slightly lower evenness (J’=0.79), than the individual sections. RUVALCABA-ORTEGA, & GONZALEZ-ROJAS & CANALES-DEL CASTILLO 257 Although the MM section did not show the highest value of diversity or evenness, nor the highest number of species or individuals per point count, its bird community shows important differences among species’ dominances. It is also the best preserved section, and the one that shows the greatest regeneration of bald cypresses at all stages. In addition, it is the only section with the Painted Bunting as a common species, which had the highest number of individuals and frequency of occurrence. This is noteworthy because the species has shown a negative population trend (1966-2006) in the Breeding Bird Surveys of U. S. (Sauer et al. 2007). On the other hand, the consequences of habitat deterioration in S and SJS sections may not have been hilly revealed yet, due to the longevity of Montezuma bald cypresses, causing a possible delay in the consequences of alteration of the riparian habitat for several decades (Rood & Mahoney 1993). The Rio Sabinas is an important site for breeding (87) and migratory (72) birds. All of the breeding species of the Rio Sabinas are confirmed (82), probable (4), or possible (1) breeders for southern Texas (south of the 31^^ parallel; Benson & Arnold 2001, Lockwood & Freeman 2004). This was expected due to the fact that these rivers have similar riparian forests dominated by bald cypress, pecan and sycamore, especially in the Plateau Region such as the Nueces, West Nueces, Frio, and South Llano Rivers (El-Hage & Moulton 2007). In northeastern Mexico, as well, there are rivers dominated by bald cypresses such as the Rio Nazas in Durango and Coahuila, and the Rio Sabinas Hidalgo and Rio Ramos in Nuevo Leon; therefore, similarities in avian communities should be expected for these riparian habitats. Twenty-six of 35 common migrant species of the riparian systems of the southwestern U.S. (Skagen et al. 2005) were found in the area, highlighting its importance as a corridor and a stopover site. It is also noteworthy that migrants from both the eastern (i.e.. Blue-headed Vireo, Yellow-throated, Pine and Black- throated Green warblers and Yellow-bellied Sapsucker) and western U.S. (i.e.. Orange-crowned and Hermit warblers, and Red-naped Sapsucker) were recorded. Finally, these data should promote management plans leading to the conservation and restoration of the riparian habitats and water flow of the Rio Sabinas especially in the disturbed sections of San Juan de Sabinas (SJS) and Sabinas (S). 258 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Acknowledgements We thank the North American Wetlands Conservation Act for the funding provided to this research project (MX-N26E1) and the Consejo Ecologico de Participacion Ciudadana de la Region Carbonifera, A. C. (CEPACI) for facilitating the logistics of the fieldwork and the contact with local communities and owners. Literature Cited American Ornithologists’ Union. 1998. Check-list of North American Birds. 7* ed. American Ornithologists’ Union. Washington, D. C., USA, 829 pp. American Ornithologists’ Union. 2000. Forty-second supplement to the American Ornithologists’ Union Check-list of North American Birds. Auk, 1 17:847-858. Anthony, R. G., G. A. Green, E. D. Forsman & S. K. Nelson. 1996. Avian abundance in riparian sections of three forest types in the Cascade Mountains, Oregon. Wilson Bulletin, 108:208-291. Arriaga, L., J. M. Espinoza, C. Aguilar, E. Martinez, L. Gomez & E. Loa (coordinadores). 2000. Regiones terrestres prioritarias de Mexico. Comision Nacional para el Conocimiento y uso de la Biodiversidad, Mexico. Available on: http://www.conabio.gob.mx/conocimiento/regionalizacion/doctos/rtp_l 52.pdf (15 Jan 2008). Banks, R. C., C. Cicero, J. L. Dunn, A. W. Kratter, P. C. Rasmussen, J. V. Remsen, Jr; J. D. Rising & D. F. Stotz. 2002. Forty-third supplement to the American Ornithologists’ Union Check-list of North American Birds. Auk, 119:897-906. Banks, R. C., C. Cicero, J. L. Dunn, A. W. Kratter, P. C. Rasmussen, J. V. Remsen, Jr; J. D. Rising & D. F. Stotz, 2003, Forty- fourth supplement to the American Ornithologists’ Union Check-list of North American Birds. Auk, 120(3):923-93 1 . Banks, R. C., C. Cicero, J. L. Dunn, A. W. Kratter, P. C. Rasmussen, J. V. Remsen, Jr; J. D. Rising & D. F. Stotz. 2004. Forty-fifth supplement to the American Ornithologists’ Union Check-list of North American Birds. Auk, 121(3):985-995. Banks, R. C., C. Cicero, J. L. Dunn, A. W. Kratter, P. C. Rasmussen, J. V. Remsen, Jr; J. D. Rising & D. F. Stotz. 2005. Forty-sixth supplement to the American Ornithologists’ Union Check-list of North American Birds. Axik, 122(3): 1026- 1031. Banks, R. C., C. Cicero, J. L. Dunn, A. W. Kratter, P. C. Rasmussen, J. V. Remsen, Jr; J. D. Rising & D. F. Stotz. 2006. Forty-seventh supplement to the American Ornithologists’ Union Check-list of North American Birds. Auk, 123(3):926-936. Banks, R. C.; R. T. Chesser; C. Cicero; J. L. Dunn; A. W. Kratter; I. J. Lovette; P. C. Rasmussen; J. V. Remsen, Jr; J. D. Rising & D. F. Stotz. 2007. Forty-eigth supplement to the American Ornithologists’ Union Check-list of North American Birds. Auk, 124(3):1109-1115. Benson, K. L. & K. A. Arnold. 2001. The Texas Breeding Bird Atlas. Texas A&M University System, College Station and Corpus Christi, TX. Accessed at: http://tbba.cbi.tamucc.edu (11 June 2008). Bray J. R. & J. T. Curtis. 1957. An ordination of the upland forest communities of Southern Wisconsin. Ecol. Monogr., 27:325-349. RUVALCABA-ORTEGA, & GONZAlEZ-ROJAS & CANALES-DEL CASTILLO 259 Colwell, R. K. 2006. Estimates 8.0: Statistical estimation of species richness and shared species from samples. Available on: http://viceroy.eeb.uconn.edu/estimates&. Colwell, R. K., C. X. Mao & J. Chang. 2004. Interpolating, extrapolating, and comparing incidence-based species accumulation curves. Ecology 85:2717-2727. DeSante, D. F. & T. L. George. 1994. Population trends in the landbirds of western North America. Studies in Avian Biology, 15:173-190. Diario Oficial de la Federacion (DOF). 6 de marzo de 2002. Norma Oficial Mexicana NOM-059-ECOL-2001, 85 pp. El-Hage, A. & D. W. Moulton. 2001. Ecologically Significant River &Stream Segments of the Region J (Plateau), Regional Water Planning Area. Texas Parks and Wildlife. Accesed at: http://tpwd.state.tx.us/publications/pwdpubs/pwd_rp_t3200_l 059d/index.phtml ( 1 1 June 2008) Finch, D. M. & W. Yong. 2000. Landbird migration in riparian habitats of the middle Rio Grande: a case study. Studies in avian biology, 20:88-98 Fisher, R. A., A. S. Corbet & C. B. Williams. 1943. The relation between the number of species and the number of in dividuals in a random sample from an animal population. Journal of Animal Ecology, 12:42-58. Flannery, M. E., S. L. Guers, T. Gardali, N. Nur & G. R. Geupel. 2004. Landbird migration at the Salton Sea: the value of desert riparian habitat. Studies in Avian Biology, 27:106-115. Howell, S. N. G. & S. Webb. 1995. A Guide to the Birds of Mexico and Northern Central America. Oxford University Press Inc., New York, EUA. 85 1 pp. Husak, M. S. & T. C. Maxwell. 1998. Golden-fronted Woodpecker (Melanerpes aurifrons). The Birds of North America Online (A. Poole, Ed.). Ithaca: Cornell Lab of Ornithology. Hutto, R. L. 1995. Can patterns of vegetation change in western Mexico explain population trends in western neotropical migrants? Pages 48-58 in M. H. Wilson, and S. A. Sader editors. Conservation of neotropical migratory birds in Mexico. Maine Agricultural and Forest Experiment Station, Misc. Publ. 727, 288 pp. lUCN. 2007. 2007 lUCN Red List of Threatened Species. Accessed at: http://www.redlist.org/ (1 March 2008). Kelly, J. F. & R. L. Hutto. 2005. On songbird migration in western North America: a case study of wood warblers. Condor, 107:197-211. Knopf, F. L. & F. B. Samson. 1994. Scale perspectives on avian diversity in western riparian ecosystems. Conservation Biology, 8:669-676. Knopf, F. L., R. R. Jonson, T. Rich, F. B. Sampson & R. C. Szaro. 1988. Conservation of riparian ecosystems in the United States. Wilson Bulletin, 100:272-284. Magurran, A. E. 2004. Measuring Biological Diversity. Blackwell Publishing, United Kingdom, 256 pp. May R. M. 1975. Patterns of species abundance and diversity. Pp. 81-120 in\ Cody M.L., Diamond J.M., eds. Ecology and evolution of communities. Cambridge, MA: Harvard University Press, 560 pp. Ohmart, R. D. 1994. The effect of human-induced changes on the avifauna of western riparian habitats. Studies in Avian Biology, 15:273-285. 260 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Periodico Oficial. 1998. 1998: Ano del “Constitucionalismo Coahuilense”. Organo del Gobiemo Constitucional del Estado Independiente, Libre y Soberano de Coahuila de Zaragoza. Tomo CV. No. 84, 45 pp. Preston, F. W. 1948. The commonness and rarity of species. Ecology, 29:254-283. Ralph, C. J., G. R. Geupel, P. Pyle, T. E. Martin, D. F. DeSante & B. Mila. 1996. Manual de metodos de campo para el monitoreo de aves terrestres. Gen. Tech. Rep. PSW-GTR- 159. Albany, CA: Pacific Southwest Research Station, Forest Service, U.S. Department of Agriculture, 44 pp. Rood, S. B. & J. M. Mahoney. 1993. River damming and riparian cottonwoods: management opportunities and problems. Pp 134-143 in B. Tellman, H. J. Cortner, and M. G. Wallace, technical coordinators. Riparian management: common threads and shared interests. U.S. Forest Service General Technical Report RM-226, Fort Collins, CO, 432 pp. Ruvalcaba-Ortega, I. & J. I. Gonzalez-Rojas. 2009. New records for Coahuila from a riparian bird community in Northern Mexico. The Southwestern Naturalist, Vol. 54 No. 4. {In press). Sauer, J. R., J. E. Hines & J. Fallon. 2007. The North American Breeding Bird Survey, Results and Analysis 1966-2006. Version 10.13.2007. USGS Patuxent Wildlife Research Center, Laurel, MD. Shannon, C. E. 1948. A mathematical theory of communication. Bell. Syst. Tech. J., 27:379-243. Skagen, S. K., C. P. Melcher, W. H. Howe & F. L. Knopf. 1998. Comparative use of riparian corridors and oases by migrating birds in southeast Arizona. Conservation Biology, 12:896-909. Sokal, R & F. J. Rohlf. 1995. Biometry. 3rd. Ed. Freeman. USA, 887 pp. Texas Parks and Wildlife Service (TPWD). 2003. Texas Threatened and Endangered Species. Texas Parks and Wildlife Department, 4 pp. Trevino-Garza, E. J., R. Lopez-Aguillon, J. Jimenez-Perez & H. Villalon-Mendoza. 2002. Plan de Manejo del Area de Proteccion de los recursos Naturales del Rio Sabinas. Gobiemo del Estado de Coahuila de Zaragoza, 219 pp. Villasenor, J. F. 2006. Habitat use and the effects of disturbance on wintering birds using riparian habitats in Sonora, Mexico. PhD Dissertation. University of Montana, 271 pp. Williams, C. B. 1964. Patterns in the balance of nature and related problems in quantitative ecology. London: Academic Press, 324 pp. IR-0 at: i.mvalcaba.o@gmail.com TEXAS J. OF SCI. 60(4):261-270 NOVEMBER, 2008 REPRODUCTION IN SEVEN SPECIES OF MICROLOPHUS (SQUAMATA: TROPIDURIDAE) FROM SOUTH AMERICA Stephen R. Goldberg Department of Biology, Whittier College Whittier, California 90608 Abstract-Information on the reproductive cycles of seven species of Microlophus (M koepckeorum, M. occipitalis, M. peruvianas, M. stolzmanni, M. theresiae, M. thoracicus and M tigris) from coastal northern South America gathered from a histological examination of gonadal material is presented. All species exhibited extended reproductive activity present in both austral spring-summer and autumn. Histological evidence is presented that M peruvianas may produce multiple clutches in the same year. Comparisons are made with the reproductive cycles of other species of Microlophus. The genus Microlophus consists of some 20 species and is restricted in distribution to South America and the Galapagos Islands (Peters & Donoso-Barros 1986). Microlophus koepcke¬ orum, M. peruvianus, M. theresiae, M. thoracicus, M. tigris are endemic to Peru (Lehr 2002). Microlophus occipitalis is known from southwestern Ecuador and northern and central Peru, and M stolzmanni is known from northwestern Peru and possibly southwestern Ecuador (Peters & Donoso Barros 1986). Information on reproduction of M peruvianus, M. theresiae, M. thoracicus and M tigris (all as Tropidurus) are in Dixon & Wright (1975) and M (as Tropidurus) occipitalis is in Watkins (1996). The purpose of this paper is to add information on the reproductive cycles of seven species of Microlophus lizards from coastal northern South America. Materials and Methods Histological examinations were performed on 313 museum specimens of seven species of Microlophus on deposit with the Natural History Museum of Los Angeles County (LACM), Los Angeles, California (Table I). Counts were made of oviductal eggs or enlarged follicles (> 4 mm length). The left testis and 262 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO 4, epididymis were removed from males; the left ovary was removed from females for histological examination. Tissues were embedded in paraffin and cut into sections at 5 pm. Slides were stained with Harris' hematoxylin followed by eosin counterstain (Presnell & Schreibman 1997). Testis slides were examined to determine the stage of the male cycle; ovary slides were examined for the presence of yolk deposition. Histological slides of most gonads from 45 juveniles: M koepckeorum (n = 4), M occipitalis {n = 6), M peruvianus {n = 3), M stolzmanni {n = 13), M theresiae (n = 6), M thoracicus (n = 8), M tigris (n = 5) were made to determine the minimum size at which maturity was reached. In cases where the gonads were extremely small, immaturity was assumed and no histology was performed. The relationship between female SVL and clutch size was examined by linear regression analysis and an unpaired t test was used to compare male and female mean body sizes (SVL) (Instat vers. 3.0b, Graphpad Software, San Diego, CA). Specimens examined. -ThQ following specimens of Microlophus were examined from the herpetology collection of the Natural History Museum of Los Angeles County (LACM). Microlophus koepckeorum {n = 46) PERU: Lambayeque Department (49083, 49084, 49086, 49087, 49089^49098, 49100, 49104, 109569, 122583-122585, 122587-122592, 122594-122606, 122608- 122613, 122615); collected 1968, 1970, 1976. Microlophus occipitalis {n = 67) PERU: La Libertad Department (48853-48861, 48866-48869, 48902-48904, 48906-48908, 122639, 125348, 136012); Lambayeque Department (48870, 48872, 48873, 48897-48901, 122617-122622, 122627-122630, 122632, 122633, 122635-122638); La Libertad Department (136013); Piura Department (48876, 48877, 48891-48895): Tumbes Department (48878-48890); collected 1968, 1976. Microlophus peruvianus (n = 45) CHILE: Antofagasta Province (122000); ECUADOR: Esmeraldas Province (154385-154387, GOLDBERG 263 154389-154394); PERU: Ancash Department (49021, 49037) Ica Department (49018, 49019, 49040, 49042, 49044-49046); La Libertad Department (136004-136011); Lima Department (9342-9345, 9347, 49023, 49024, 49026, 49028-49030, 49032, 49034, 49036, 49050, 49055-49057); collected 1965, 1966, 1968, 1984. Microlophus stolzmanni (n= 50) PERU: Cajamarca Department (49107, 49108, 49110-49116, 49119-49123, 49126-49129, 49131, 49132, 122641, 122646, 122649-122651, 122653-122657, 122659, 122660, 122662-122667, 122669, 122670, 122672-122676, 122678- 122682); collected 1968, 1976. Microlophus theresiae {n = 29) PERU: Ica Department (49060- 49063, 49075-49077); Lima Department (LACM 49064, 49068-49072, 49074, 49079, 122692-122698, 122700-122706); collected 1968, 1976. Microlophus thoracicus (^ = 50) PERU: Ancash Department (48911, 48914-48918, 48953, 48954, 48956-48963, 48965-48967, 48969-48974, 48976-48979, 122737, 122738, 122742, 122746, 122747); Ica Department (122726-122731, 122733-122736); La Libertad Department (48951); Lambayeque Department (48922, 48924); Lima Department (48980, 48983, 48984); collected 1968, 1976. Microlophus tigris (n = 26) PERU: Lima Department (48830- 48838, 48841-48847, 48849, 122712-122714, 122716, 122718-122722); collected 1968, 1976. Results And Discussion Microlophus koepckeorum -The mean SVL of males was signi¬ ficantly larger than that of females (Table 1). All 21 males examined (May =10, June = 2, August = 1, November = 8) were undergoing spermiogenesis (= sperm formation). In this stage, the seminiferous tubules were lined by spermatozoa or groups of metamorphosing spermatids and the epididymides contained sperm. The smallest reproductively active male (122592) was undergoing spermiogenesis, measured 52 mm SVL and was from May. 264 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO 4, Table 1. Sample sizes (w), mean sizes (snout- vent length, mm) ± SD and range for males of seven species of Microlophus from South America. *Indicates mean snout-vent length of males was significantly larger than that of females (unpaired ^-test). DF t P Males Females n X±SD (Range) n X±SD (Range) M. koepckeorum* 40 6.9 <0.001 21 67.6±6.8 (52-78) 21 55.7±4.1 (50-67) M. occipitalis* 59 4.9 <0.001 28 62.8±11.9 (35-79) 33 51.5±5.1 (44-65) M. peruvianas* 40 2.9 <0.001 17 86.8±18.8 (50-109) 25 73.0±12.6 (55-93) M. stolzmanni* 35 6.1 <0.001 19 88.2±17.4 (65-120) 18 62.1±5.3 (53-73) M. theresiae* 21 4.7 0.0001 6 68.0±3.9 (62-72) 17 59.8±3.6 (53-67) M. thoracicus* 40 4.7 <0.0001 23 63.2±7.8 (43-77) 19 53.9±4.3 (47-64) M. tigris - - - 16 61.9±13.8 (40-80) 5 55.0±8.5 (45-67) Monthly stages in the ovarian cycle are in Table 2. Reproductively active females were found in May- June and November. Mean clutch size {n = 13) was 3.2 ± 0.69 SD, range = 2-5. Linear regression analysis revealed a significant positive correlation between female body size (SVL) in mm and clutch size for 13 M koepckeorum females (r = 0.82, P = 0.0007). The smallest reproductively active females (both from May) each measured 50 mm SVL (122595, early yolk deposition and 122601, three follicles > 4 mm). Dixon & Wright (1975) reported M koepckeorum mature at 50 mm SVL, and reproductively active females were found in August, November and December with 2-4 yolked ovarian follicles or oviductal eggs. Microlophus occipitalis -JhQ mean SVL of males was signifi¬ cantly larger than that of females (Table 1). All 28 males examined (May = 6, June = 2, November = 7, December = 13) were undergoing spermiogenesis. The smallest reproductively active male (spermiogenesis) measured 35 mm SVL (48853) and was from November. Reproductively active females were found June, August, November and December (Table 2). The smallest reproductively active female (48906, two follicles > 4 mm, measured 44 mm SVL and was from December). Mean value for nine clutches was 4.5 ±1.8 SD, range = 2-8. Linear regression GOLDBERG 265 Table 2. Monthly stages in ovarian cycles of seven species of Microlophus from Peru. Month n No yolk deposition Early yolk deposition Follicles > 4 mm Oviductal eggs M koepckeorum May 12 2 1 5 4 Jun 5 0 2 1 2 Nov 4 0 2 1 1 M. occipitalis Jun 9 2 2 4 1 Aug 1 0 1 0 0 Nov 7 3 0 3 1 Dec 16 15 0 1 0 M. peruvianus May 4 1 * 1 0 2 Jun 2 0 0 2 0 Aug 1 0 0 1 0 Oct 1 1 0 0 0 Nov 10 4** 1 0 5 Dec 7 2** 2 2 1 M. stolzmanni Jun 13 0 1 10 2 Nov 5 0 0 3 2 M. theresiae Jun 11 0 2 9 0 Dec 6 5 0 0 1 M. thoracicus May 5 1 1 2 1 Dec 14 2 0 4 8 M. tigris May 1 1 0 0 0 Dec 4 2 0 1 1 = one M. peruvianus female from May contained corpora lutea from a previous clutch and concomitant yolk deposition for a subsequent clutch; = one M. peruvianus female from November and another from December each contained corpora lutea but were not undergoing yolk deposition. analysis revealed a significant positive correlation between female body size (SVL) in mm and clutch size for 9 M occipitalis females (r = 0.75, P = 0.021). Dixon & Wright (1975) reported oviductal eggs in females from July, August and November, clutches of 2-4 eggs were produced. Males mature between 50-55 mm SVL and females between 45-47 mm SVL (Dixon & Wright 1975). Watkins (1996) studied a population of M occipitalis at Guayas Province, 266 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO 4, Ecuador. He reported males (mean SVL = 63.0 ± 15.7 mm) were larger than females (mean SVL = 55.1 ± 7.5 mm) and the smallest female with oviductal eggs or enlarged follicles measured 48 mm SVL. Burt (1935) reported that a series of 15 M occipitalis from Ecuador collected in January exhibited gradation in size from juveniles to adults suggesting an extended breeding season. Microlophus peruvianus -Tht mean SVL of males was signifi¬ cantly larger than that of females (Table 1). Sixteen of 17 males examined were undergoing spermiogenesis (May = 5, June = 6, November = 4, December =1). One November male (SVL = 66 mm) appeared to have completed spermiogenesis. The germinal epithelium was reduced to a few layers in thickness, occasional metamorphosing spermatids but no spermatozoa were present. The smallest reproductively active male (spermiogenesis) measured 63 mm SVL (49021) and was from November. Reproductively active females were present May, June, August and November (Table 2). The smallest reproductively active M peruvianus female contained two oviductal eggs (49028) measured 56 mm SVL and was from November. Mean clutch size for 13 gravid females was 3.5 ± 1.3 SD, range = 2-6). Linear regression analysis revealed a significant positive correlation between female body size (SVL) in mm and clutch size for 13 M peruvianus females (r = 0.60, P = 0.032). Dixon & Wright (1975) reported M peruvianus females were reproductively active during July- September and November and December. One M peruvianus female from May (154389) contained corpora lutea from a previous clutch and concomitant vitellogenic follicles for a subsequent clutch indicating M peruvianus can produce multiple clutches in the same year. One female each from November (49032) and December (49055) contained corpora lutea indicating eggs had recently been deposited. There was no concomitant yolk deposition for a subsequent clutch. Leyton et al. (1981) reported males and females of M (as Tropidurus) peruvianus were reproductively active throughout the year in northern Chile. GOLDBERG 267 Microlophus stolzmanm-Th.Q mean SVL of males was signifi¬ cantly larger than that of females (Table 1). All 19 males examined were undergoing spermiogenesis (May = 1, June = 8, November = 10). The smallest reproductively active male (spermiogenesis) measured 65 mm SVL (122672) and was from June. Reproductively active females were found in June and November (Table 2). The smallest reproductively active female (49128, measured 53 mm SVL) and was from November. Mean clutch size for 17 females was 2.6 ± 0.86, range = N4. Linear regression analysis revealed a significant positive correlation between female body size (SVL) in mm and clutch size for 13 M stolzmanni females (r = 0.53, P = 0.029). Dixon & Wright (1975) reported M stolzmanni females were reproductively active in November, matured between 65-76 mm SVL and produced 3-4 oviductal eggs. Microlophus theresiae.-ThQ mean SVL of males was signifi¬ cantly larger than that of females (Table 1). All 6 males examined were undergoing spermiogenesis (June = 3, November = 1, December = 2). The smallest reproductively active male (spermiogenesis) measured 62 mm SVL (49072) and was from December. Reproductively active females were found in June and December (Table 2). Mean clutch size for 10 females was 3.4 ± 0.84, range = 2-5. Linear regression analysis revealed no significant positive correlation between female body size (SVL) in mm and clutch size for 10 M theresiae females. The smallest reproductively active female (122696, 3 follicles > 4 mm), measured 58 mm SVL and was from June. Dixon & Wright (1975) reported M theresiae females contained oviductal eggs or enlarged follicles in December. Microlophus thoracicus mean SVL of males was signifi¬ cantly larger than that of females (Table 1). Seventeen of nineteen males were undergoing spermiogenesis: (May = 3, November = 6, December = 8). One male from May was undergoing late recrudescence (renewal of germinal epithelium for the next period of spermiogenesis). A few metamorphosing spermatids were noted. 268 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO 4, Another May male was in late spermiogenesis, clusters of metamorphosing spermatids and spermatozoa were present. The germinal epithelium was reduced in cellularity. The smallest reproductively active male (spermiogenesis) measured 43 mm SVL (48983) and was from December. Reproductively active females were found in May and December (Table 2), The smallest reproductively active female (48966, 2 oviductal eggs) measured 46 mm SVL and was from December. Mean clutch size for 15 females was 2.5 ± 0.83, range = 1-4. Linear regression analysis revealed a significant positive correlation between female body size (SVL) in mm and clutch size for 15 M thoracicus females (r = 0.60, P = 0.018). Dixon & Wright (1975) reported M thoracicus females were reproductively active in November and December, mature at 45-50 mm SVL and produce 1-5 eggs per clutch with a mean of 3.1 eggs for 40 females. Microlophus Z/grA.-There was no significant difference between mean males and females in SVL (Table 1). Fourteen of 16 males examined were undergoing spermiogenesis: (May = 8, December = 6). Two males from December were in recrudescence prior to the next period of sperm formation. Germinal epithelium contained sufficient numbers of cells to support spermiogenesis, A few metamorphosing spermatids but, as yet, no spermatozoa were present. The smallest reproductively active male (spermiogenesis) measured 40 mm SVL (122713) and was from May. Reproductively active females were found in December (Table 2). The smallest reproductively active female (48834, 2 follicles > 4 mm) measured 56 mm SVL and was from December. Mean clutch value for two females was 3.0 ± 1.4 SD, range = 2-4. There was insufficient data to perform regression analysis to examine the relation between body size and clutch size. Dixon & Wright (1975) reported M tigris females were reproductively active in September, November and December and produced clutches of 2-5 eggs. It appears that other species of Microlophus from Peru and elsewhere have extended reproductive cycles. Goldberg & Rodriguez (1986) reported on reproduction in M quadrivittatus and GOLDBERG 269 M theresioides (both as Tropidurus) from northern Chile. In both species maximum numbers of males were undergoing spermiogenesis in spring although varying numbers of spermiogenic males were found in most months indicating an extended reproductive cycle. Some males with regressed testes were found in summer; female reproductive activity (enlarged follicles or oviductal eggs) was limited to spring (Goldberg & Rodriguez 1986). This contrasts with six of the seven species of Microlophus presented herein which were reproductively active in both austral winter and spring (spring female data was limited to only one specimen of M tigris). Furthermore, Vitt & Goldberg (1983) similarly reported some males of Microlophus (as Tropidurus) torquatus from northeastern Brazil were undergoing spermiogenesis and females were reproductively active in all months. Subsequent examination of the reproductive biology of other Microlophus species are needed before the diversity of reproduction exhibited by these lizards can be ascertained. Acknowledgments I thank Christine Thacker for permission to examine Microlophus sp. Literature Cited Burt, C. E. 1935. Notes on a collection of lizards from western Mexico and tropical America. Trans. Amer. Micros. Soc., 54:167-178. Dixon, J. R. & J. W. Wright. 1975. A review of the lizards of the iguanid genus Tropidurus in Peru. Nat. Hist. Mus. Los Angeles Co, Contrib. Sci., 271:1-39 pp. Goldberg, S. R. & E. Rodriguez. 1986. Reproductive cycles of two iguanid lizards from northern Chile, Tropidurus quadrivittatus and Tropidurus theresioides. J. Arid Environ., 10:147-151. Lehr, E. 2002. Amphibien und Reptilien in Peru. Natur und Tier, Munster. 206 pp. Leyton, V., A. Veloso & E. Bustos. 1981. Modalidad reproductiva y actividad ciclica gonadal en lagartos iguanidos de distinctos pisos altitudinales del interior de Arica (Lat. 18°10’S). Pp. 293-316 in Vol. I La vegetacion y los vertebrados inferiores de los pisos altitudinales entre Arica y el Lago Chungara (A. Veloso & E. Bustos, eds.). Proyecto MAB-6, UNEP-UNESCO-1 105-77-01. 270 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO 4, Peters, J. A. & R. Donoso-Barros. 1986. Catalogue of the Neotropical Squamata: Part II Lizards and Amphisbaenians. Smithsonian Institution Press, Washington, D.C. 293 pp. Presnell, J. K. & M. P. Schreibman. 1997. Humason’s Animal Tisssue Techniques, 5* Ed. The Johns Hopkins University Press, Baltimore, 572 pp. Vitt, L. J. & S.R. Goldberg. 1983. Reproductive ecology of two tropical iquanid lizards: Tropidurus torquatus and Platynotus semitaeniatus. Copeia, 1983:131- 141. Watkins, G. G. 1996. Proximate causes of sexual size dimorphism in the iguanian lizard Microlophus occipitalis. Ecology, 77:1473-1482. SRG at; sgoldberg@whittier.edu TEXAS T OF SCL 60(4):27E280 NOVEMBER, 2008 NOTES ON FOOD ATTRACTION AND THE DEMOGRAPHICS OF BLUE CRAB (CALLINECTES SAPIDUS) TRAPPING IN TRES PALACIOS BAY, TEXAS Joel D. Anderson and Dusty L* McDonald Perry R. Bass Marine Fisheries Research Station Texas Parks and Wildlife Department Palacios Texas 77465 Abstract.-The null hypothesis that blue crabs which enter typical commercial- style baited traps are random draws from populations with equal sex ratios and physical characteristics was evaluated. Trap demographic data (sex, size and number of crabs) were collected during ten single-day trials comprising six traps each {n = 60) over a 3-week period in April 2006, These data indicate that blue crabs tend to be caught in same-sex assemblages more frequently than expected by chance, and that males caught in crab traps have different width- weight ratio trajectories than do females. Interestingly, blue crabs in Tres Palacios Bay are more readily attracted to remains of fish that are endemic and common in the Gulf of Mexico, rather than those that are exotic and/or relatively rare. These data relate directly to previously unexplored aspects of the blue crab fishery in Texas and therefore will be usefiil to fishery managers. Blue crabs (Callinectes sapidus Rathbun) are subject to a significant commercial fishery on the coast of Texas, harvested at a rate surpassed only by shrimp and oysters (in total organisms landed) (Suttton & Wagner 2007). Landings have declined considerably since 1987, due to a combination of many factors including overharvest (Sutton & Wagner 2007). Currently, Texas Parks and Wildlife (TPW) regulates the harvest of blue crabs by a minimum-size limitation of five inches (127 mm), a ban on harvest of egg-bearing females, commercial trap limitations (200 traps) and design specifications, and a 10-day to 30-day closure of the fishery each year (February or March). To evaluate trends in blue crab abundance and biomass, TPW typically relies on fishery- independent data collected during annual routine monitoring. However, sparse data is available that pertains to the demographics of crabs specifically targeted by current trapping methodologies. Although abundant data is available detailing the effects of trap design on retention of various crab size ranges (Guillory & Prejean 272 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 1997; Guillory 1998; Guillory & Hein 1998a; 1998b) there is a lack of available data on the size and sex ratios of crabs typically retained in traps in the western Gulf of Mexico. Behavioral and ecological characteristics such as mutual antagonism during foraging (Mansour & Lipcius 1991; Clark et al. 1999) and intra- population habitat partitioning (Hines et al. 1987) may result in different demographic characteristics between wild crab populations, and what is typically found in commercial traps. Here, the demographics of crabs captured in commercial-style traps in Tres Palacios Bay, Texas are described in an effort to improve the data upon which future management decisions are based. Methods Six identical commercial- style crab traps which consisted of a coated wire frame throughout, a bait compartment and four entrance funnels were used in this study. The dimensions of the traps were 60 cm width, by 60 cm length, by 44 cm height. Entrance funnels were approximately oval in shape, were spaced evenly on each of the four sides of the trap and were 26 cm wide by 12 cm high. The bait compartment was located central to the bottom of the trap, and was 22 cm in height by 12 cm in diameter. While commercial traps in Texas are required to include 6.03 cm escape vents in each retaining chamber, traps used in experimental trials contained no escape vents and theoretically retained all crabs captured during the duration of the trial. Traps were deployed in a 2 by 3 grid system adjacent to a rock jetty (Fig. 1). The first row of traps was located approximately 100 m from the shoreline, with each additional row located 20 m apart. In each row, traps were located 20 m and 40 m from the face of the jetty. During the first three weeks of April 2006, traps were baited and set between 0730 h and 0800 h on each day of the trial. Trials were conducted Mon-Thurs of the first two weeks and Mon-Tue of the final week, for a total of ten trials. Bait was prepared by measuring an equal amount of cut bait menhaden (200g) into the bait compartment of each trap. Three species of menhaden were used ANDERSON & MCDONALD 273 for bait; finescale menhaden (Brevoortia gunteri), Gulf menhaden {B. pair onus) and Atlantic menhaden (B. tyr annus). Bait was distributed into traps in randomized design each day in order to test for differences in attraction among bait species. Traps were emptied and crabs collected following 24 h of exposure. Traps were checked as quickly as possible by two researchers in order to limit the amount of time between traps. Hydrological data were recorded prior to checking traps. Hydrological data consisted of water temperature (°C), salinity (practical salinity units, or psu) and dissolved oxygen (mg/L). Tidal data for each day of the three- week trial was obtained from the National Oceanic and Atmospheric Administration’s online tidal database (NOAA 2007), and included high tide and low tide measured as the difference between the recorded water level and the mean water level on site. The difference between high and low tide for each day of the trial was calculated and used as a proxy for the volume of water movement into and out of the site. Crabs were extracted from the sampling area without return for the duration of the study. Each crab was weighed to the nearest g, and the greatest width of the carapace in mm was recorded. Sex was determined by the shape of the abdomen ventrally. In the case of escape, crabs were recorded as “lost” and were used only for demographic count data. In addition to blue crabs, the occurrence of other species, such as hermit crabs and (rarely) fishes were recorded by species, but no measurements were taken and these organisms were released back into the study site. The assumption that the sex ratio of crabs caught in single traps is likely to be representative of the sex ratio in the population was tested by constructing contingency tables with single-sex and multiple-sex trap counts (rows) by the number of crabs caught within a specified trap (columns, 72 = 2, 3, 4 and 5-crab traps). The expected frequency of single-sex traps for each treatment was calculated using the estimated frequency of males and females in the population. For instance, the expected frequency of traps 274 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 containing two crabs of the same sex was calculated as {n (p^ + q^)), where n = the overall number of traps containing two crabs, p = the overall frequency of males in the study and q = the overall frequency of females. This value was then subtracted from 1.0 to obtain the expected frequency of traps containing both sexes. Similar expected frequencies were obtained for traps containing three, four and five crabs. Cell values for traps which collected more than three crabs were combined into a single treatment to minimize the effects of small expected frequencies as described by Cochran (1952). Observed and expected frequencies of same and multi-sex traps were compared using a test of homogeneity, and significance was assessed via comparison to the null distribution with an arbitrary value of a = 0.05 for all significance tests. The relationship between crab width and total weight was assessed via the general linear model, as implemented in SAS (version 8.02, SAS Inst., Inc., Cary, NC). Weight was treated as the response variable, and the predictor variables considered in the model were sex, width and sex/width. Sex was included as a predictor after observing what appeared to be different trajectories of the width-weight curves of males and females. Results Through the course of the study, salinity (range 30.0-31.4 psu), and dissolved oxygen (5.8 - 6.9 mg/L) remained relatively con¬ sistent at the study site. Female crabs tend to be found in higher salinity than males (Tagatz 1971, and references therein), and more females than males were captured in this study. However, because of the relative consistency of high salinity throughout all trials, the effects of variability in salinity on crab sex ratios could not be evaluated. Temperature also remained relatively consistent among trials (18.9 - 24.5 °C) although fewer crabs were captured at the lowest temperature trial than any other trial. Otherwise, there was not a significant relationship between the number of crabs captured and either temperature = 0.075, p = 0.446) or tide {/ = 0.017, p = 0.737). ANDERSON & MCDONALD 275 Shoreline Figure 1. Study design of Callinectes sapidus study. Traps are labeled A-F and were located at indicated distances from the shoreline and jetty. A total of 86 crabs were captured through the duration of the study, including 50 females, 35 males and a single individual that was of undetermined gender prior to escape. Overall trap success throughout the study ranged from 12 crabs in trap B (Fig. 1) to 20 crabs caught in trap E. Heterogeneity in success among trap positions was not significantly different from the distribution expected under equal success {jC = 3.02, p = 0.696), suggesting that crabs entered each trap with equal probability. However, there was a significant difference in success between bait types (;f = 7.19, j!? = 0.027). Whereas crabs entered traps baited with B. tyrannus at approximately the rate (n = 26) expected under equal distribution (equal distribution = 86 crabs/3 bait types, or 28.7 crabs per bait), traps baited with B. pair onus were visited more frequently (n = 38), and traps baited with B. gunteri less frequently (n = 22), than would be expected under equal distribution. Both the total number of positive traps (traps capturing at least one crab) and the total number of crabs captured were highest in traps baited with B. patronus (Fig. 2). 276 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Figure 2. The trap success of each of three species of crab bait, Gulf menhaden Brevoortia patronus, Atlantic menhaden B. tyrannus and fmescale menhaden B. gunteri. The number of positive traps (traps with at least one crab) and overall number of crabs per bait are counted on the left axis, whereas the trap density (crabs/trap) is indicated on the right axis. Crabs tended to be captured in same-sex assemblages more frequently than would be expected assuming entrance at rates proportionate to the local sex ratio, calculated from the expected ratio of females to males over all samples = 7.89, p = 0.005). For instance, the frequency of females in the study was 0.588, and the frequency of males was 0.412; however, of the two traps that captured four crabs, each contained only females. Similarly, three of five 3-crab traps contained single-sex assemblages. The width- weight relationship trajectory was also different between male and female crabs (p = 0.004). Much of the variance in weight could be accounted for by differences in width (F = 193.51, < 0.001), as compared to sex (F = 30.59, p < 0.001). However, males tended to be heavier than females of equal width, especially in larger (and presumably, older) individuals (Fig. 3). In addition to blue crabs, other species were captured during the duration of the study, including stiped hermit crabs {Clibanarius vittatus, n = 82), hardhead catfish (Arius felis, n 6) and a single red drum (Sciaenops ocellatus). While the effect of these species ANDERSON & MCDONALD 111 Figure 3. Width- weight relationship of male (open circles) and female (closed boxes) blue crabs, CalUnectes sapidus. on the entrance of blue crabs into traps was untested, the capture of these species after only a single night of trap placement has implications for ghost fishing (Guillory 1993) with commercial- style blue crab traps. Discussion In previous studies, the number and size of crabs entering commercial-style traps was shown to vary seasonally (Guillory 1993). However, to the author’s knowledge, sex-ratios within individual traps have not previously been examined. Here, blue crabs tended to be captured in same-sex groups more often than would be expected by chance. While this result seems extraordinary, the finding is not without biological basis. For instance, Hines et al. (1987) demonstrated significant habitat partitioning by size, molt stage and sex. Broad movements of both sexes, but females in particular, are affected primarily by reproductive cycles and result in significant differences in resource usage. As a result, it is not surprising to find that males and females may be collected in same-sex assemblages. These data also indicate divergent width- weight trajectories for males and females. The finding that males tend to be heavier than females of equal width was first reported by Newcombe et al. (1949) and has since 278 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 been corroborated by numerous authors working in Texas (Pullen & Trent 1970), South Carolina (Olmi & Bishop 1983), Florida (Tagatz 1965) and even Turkey (Atar & Secer 2003). If weight can be used as an indication of crab maturity, this finding indicates that mature males of market size will be encountered less often than females in baited traps. However, in the duration of the current study, only four crabs were encountered that were smaller than market size; of these, two were male and two were female. Thus, if these numbers are reflective of the fishery as a whole, it is likely that crabs are harvested equally regardless of gender. Additionally, Olmi & Bishop (1983) reported that in addition to sex and maturity, molt stage was a considerable source of variance in weight of individual crabs. Thus it seems unlikely that size requirements for com¬ mercially captured crabs in Texas result in biased exploitation of either male or female crabs, although this finding could potentially be reevaluated with fishery-dependent data. The finding that blue crabs were statistically more likely to be caught in traps baited with Gulf menhaden B. patronus is very significant. The Gulf menhaden is one of the more common fishes in the western Gulf of Mexico and census estimates for this species suggest that the Gulf population may number in the hundreds of millions (Avise et al. 1989) or more. In contrast, the finescale menhaden B. gunteri exist in smaller population sizes and are encountered much more infrequently (Dahlberg 1970, Anderson & McDonald 2007), whereas the Atlantic menhaden B, tyrannus is generally not encountered in the Gulf. Thus, the implication is that blue crabs in Tres Palacios Bay exploit species which are endemic and common in the Gulf of Mexico, rather than species that are exotic or rare. Blue crabs rely heavily upon olfactory-mediated chemical cues for foraging (Weissburg & Zimmer-Fuast 1994, Keller & Weissburg 2004), and thus may be more readily attracted to cues which are frequently encountered. However, this result must be tempered by at least one major caveat. Both Gulf and Atlantic “large-scaled” menhaden species tend to have softer flesh and are oilier than their small-scaled menhaden counterparts (such ANDERSON & MCDONALD 279 as the finescale menhaden, Dahlberg 1970). The authors are in agreement that rotten tissue from finescale menhaden tends to be much less malodorous than either of the large-scaled baits, although this was not quantified scientifically. Nevertheless, the difference in attraction noted here may be simply attributed to the strength of odor plumes caused by physiological differences between species. Behavioral studies examining bait preference in a closed system would be helpfijl but were not conducted. Irrespective of the cause, the finding of differences between bait effectiveness is significant for the following reason. During the course of this study, the authors purchased a box of “crab bah” from a local fish dealer, and were given a box of fish that was primarily (> 99%, by weight) menhaden. Using previously described molecular and morphological characters (Anderson 2007, Anderson & McDonald 2007), it was determined that the box was filled with Atlantic menhaden, rather than the local (and presumably more effective) Gulf menhaden. Literature Cited Anderson, J. D & D. L. McDonald. 2007. Morphological and genetic investigations of two western Gulf of Mexico menhadens (Brevoortia spp,). J. Fish Biol. 70(sa):139-147. Anderson, J. D. 2007. Systematics of the North American menhadens: molecular evolutionary reconstructions in the genus Brevoortia (Clupeiformes: Clupeidae). Fish Bull, 71:235-249. Atar, H. H. & S. Se^er. 2003, Width/length- weight relationships of the blue crab {CalUnectes sapidus Rathbun 1896) population living in Beymelek lagoon lake. Turk. J. Vet. Anim, Sci., 27:443-447. Avise, J. C., B. W. Bowen & T. Lamb. 1989. DNA Fingerprints from hypervariable mitochondrial genotypes. Mol. Biol. EvoL, 6(3):258-269. Clark, M. E., T. G. Walcott, D. L. Wolcott & A. H. Hines. 1999. Intraspecific interference among foraging blue crabs CalUnectes sapidus: interactive effects of predator density and prey patch distribution. Mar. Ecol. Prog, Ser., 178:69-78. Cochran, W. G. 1952. The chi-square goodness-of-fit test. Arm. Math. Stat., 23:315-345. Dahlberg, D. W. 1970. Atlantic and Gulf of Mexico menhadens, genus Brevoortia (Pisces: Clupeidae). Bull. Fla. Mus. Nat. Hist. Biol. Sci., 15(3):91-162. Guillory, V. 1993. Ghost fishing by blue crab traps. N. Am. J. Fish. Manag., 13:459..466. 280 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Guillory, V. 1998. Blue crab, Callinectes sapidus, retention rates in different trap meshes. Mar. Fish Rev., 60(l):35-37. Guillory, V. & S. Hein. 1998a. A review and evaluation of escape rings in blue crab traps. J. Shellfish Res., 17(2):551-559. Guillory, V. & S. Hein. 1998b. An evaluation of square and hexagonal mesh blue crab traps with and without escape rings.. J. Shellfish Res., 17(2):561-562. Guillory, V. & P. Prejean, 1997. Blue crab, Callinectes sapidus, trap selectivity studies: mesh size. Mar. Fish Rev., 59(1):29-31 Hines, A. H., R. N. Lipcius & A. M. Haddon. 1987. Population dynamics and habitat partitioning by size, sex and molt stage of blue crabs Callinectes sapidus in a subestuary of central Chesapeake Bay. Mar. Ecol. Prog. Ser., 36:55-64. Keller, T. A. & M, J. Weissburg. 2004. Effects of odor flux and pulse rate on chemosensory tracking in turbulent odor plumes by the blue crab, Callinectes sapidus. Biol. Bull., 207:44-55. Mansour, R. A. & R. N. Lipcius. 1991. Density-dependent foraging and mutual interference in blue crabs preying upon infaunal clams. Mar. Ecol. Prog. Ser., 72:239-246. Newcombe, C. L., Campbell, F. & A. M. Eckstine. 1949. A study of the form and growth of the blue crab Callinectes sapidus Rathbum. Growth, 13:71-96. NOAA 2007. National Oceanic and Atmospheric Association online tidal database; http://www.tidesonline.nos.noaa.gov: last accessed June 2007. Olmi, E. J. Ill & J. M. Bishop. 1983. Variations in total width- weight relationships of blue crabs, Callinectes sapidus, in relation to sex, maturity, molt stage and carapace form. J. Crustac. Biol., 3(4):575-581. Pullen, E. J. & W. L. Trent. 1970. Carapace width-total weight relation of blue crabs from Galveston Bay, Texas. Trans. Amer. Fish. Soc., 99: 795-798. Sutton, G. & T. Wagner. 2007. Stock assessment of blue crab {Callinectes sapidus) in Texas coastal waters. Management Data Series No. 249, Texas Parks and Wildlife, Austin, Texas, 46 pp. Tagatz, M. E. 1965. The fishery for blue crabs in the St. Johns River, Florida, with special reference to fluctuation in yield between 1961 and 1962. USFWS Special Scientific Report — Fisheries, No. 501. Washington, DC, 1 1 pp. Tagatz, M. E. 1971. Osmoregulatory ability of blue crabs in different temperature- salinity combinations. Chesapeake Sci., 12:14-17. Weissburg, M. J. & R. K. Zimmer-Faust. 1994. Odor plumes and how blue crabs use them in finding prey. J. Exp. Biol., 197:349-375. JDA at: Joel.Anderson@tpwd.state.tx.us TEXAS J= OF SCL 60(4):28E298 NOVEMBER, 2008 SPATIAL AND TEMPORAL DISTRIBUTIONS OF PLANKTONIC DIATOMS IN A SUBTROPICAL BAYOU, ALONG THE UPPER TEXAS COAST Antonietta Quigg and Linda Roehrborn Department of Marine Biology Texas A&M University at Galveston 5007 Avenue U Galveston, Texas, 77551 and Department of Oceanography, Texas A&M University, 3146 TAMU College Station, Texas, 77843 Abstract.-A study of planktonic diatom (Bacillariophyceae) community spatial and temporal distributions was carried out in Offatts Bayou, a small embayment within the larger Galveston Bay complex in southeast Texas. The dominant diatom genera were Chaetoceros, Ditylum, Rhizosolenia, Coscinodiscus, Guinardia, Dactyliosolen, Odontella and Lithodesmium, in this order. Over 20 other diatom genera were represented at different times including some known to produce toxins (e.g,, Pseudonitzschia), while others only appeared in surface waters after strong wind induced mixing events (e.g., Navicula). The spatial and temporal patterns observed for the dominant planktonic diatoms followed changes in salinity and temperature. Other environmental variables (light, nutrients, grazing) were not measured as part of this initial investigation but may have been important. Mean Chaetoceros, Ditylum, Odontella and Lithodesmium standing crops were highest at the open end of the bayou, while Guinardia and Dactyliosolen numbers were highest in the most flow-restricted regions of the bayou. Spatial distributions of Rhizo¬ solenia and Coscinodiscus were homogenous throughout Offatts Bayou. Understanding natural diatom succession patterns may provide important insights into normal variations in community composition. Diatoms are potentially a powerful biomonitoring tool for future assessments of the impacts of eutrophication, climate change and/or human induced ecosystem dysfunction. Climate change and sea level rise will pose complex effects on phytoplankton communities, particularly those in coastal environ¬ ments. Eutrophication induced decline in ecosystem fimction in coastal areas has resulted in algal blooms, red and/or brown tides (Hallegraeff 1993; Nixon 1995; Howarth et al. 2000), fish disease, fish kills, and the development of hypoxic areas (Paerl et al. 1998; Thronson & Quigg 2008). Like bays and estuaries, coastal bayous are especially susceptible to contamination because they have intrinsically low flushing rates and limited freshwater inflow. In their tidal reaches, bayous are high in suspended sediments and dissolved organic carbon, and low in dissolved oxygen, especially 282 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 from late spring to early fall (e.g., Dickinson Bayou, Texas; Quigg et al. 2009). Given that diatoms (Division Bacillariophyta) are important primary producers in coastal waters, making up a large fraction of the phytoplankton community in many places (Omolfsdottir et al. 2004a; Omolfsdottir et al. 2004b; Bukyates & Roelke 2005; Lavoie et al. 2006), changes in their dynamics may be used as a sensitive biomonitor of the influence of the above processes on ecosystem function. Diatoms respond quickly to physical, chemical and biological perturbations (e.g., Omolfsdottir et al. 2004a; Omolfsdottir et al. 2004b). Yet few studies have examined diatom community composition and succession in coastal systems (Resende et al. 2005; Cetinic et al. 2006) despite growing concerns for ecosystem health in these, the most densely populated, developed and growing areas in the world. Galveston Bay (Fig. 1) is the second largest estuary (ca. 1550 km ) on the Texas coast. It faces conservation issues due to high density industrialization and urbanization throughout its watershed (GBEP 2001; TWDB 2007). A human population of four million live within its five bordering counties; this figure doubles when considering the entire watershed which encompasses two of the largest metroplexes in Texas: Houston and Dallas-Fort Worth (TWDB 2007). Galveston Bay is the most productive of all Texas’ estuaries with an oyster production that is unsurpassed in the U.S., and a recreational fishery that is worth millions of dollars annually (GBEP 2001; Lester & Gonzalez 2002; TWDB 2007). Phytoplankton biomass (measured as chlorophyll a) has been declining in Galveston Bay since the 1970’s (Lester & Gonzalez 2002). The bay now sustains a moderate to low phytoplankton biomass of 4 to 15 ug chlorophyll a L’^ (Santschi 1995; Quigg et al. 2007). The decrease is thought to be a response of the bay to improved water quality as a result of the Clean Water Act of 1970 (H-GAC 2006). Concurrent decreases in nitrogen and phosphate loading have also been recorded (Santschi 1995; Lester & Gonzalez QUIGG & ROEHRBORN 283 29.296 29.280 29.264 Figure L Galveston Bay is the largest estuary along the Texas coast. Offatts Bayou is a small embayment within this complex. Sampling site locations for water collected in the study area are indicated with filled circles. 2002) along with increased water clarity (Lester & Gonzalez 2002). Although phytoplankton blooms occur at different times of the year in the bay, they typically do not reach proportions which may be considered 'harmful’. The appearance of the diatom Nitzchia pungens forma multiseries in 1989 in Galveston Bay (Fryxell et al. 1990) and in 1990 in Offatts Bayou (Reap 1991) first raised concerns for human health and for the local fisheries nearly two decades ago. The presence of this and other diatoms (and species 284 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 of other phytoplankton) capable of producing toxins remains an issue of concern for scientists, resource managers and residents. In order to develop an effective biomonitoring tool, it must be simple, fast, inexpensive, representative and reliably measured. Planktonic centric diatoms potentially fulfill all these criteria in that they are easily collected, readily identifiable (at least to generic level), and cosmopolitan. Students, volunteers and state agency personal can easily participate in a biomonitoring program involving diatoms. Phytoplankton other than diatoms are often more difficult to identify without sufficient training or molecular methods; some are very fragile and/or sensitive and may be broken by improper handling and/or preservation techniques. Smaller phytoplankton species such as cyanobacteria are often missed by nets or require specialized collection techniques. Filamentous species require certain assumptions in the enumeration process and often form clumps or masses, complicating the process fiarther. Phytoplankton that typically do not occur in surface waters also require specialized collection techniques. Hence, planktonic centric diatoms were selected as a possible biomonitoring tool. This study reports the findings of a year-long study which examined patterns of diatom {Chaetoceros, Ditylum, Rhizosolenia, Coscinodiscus, Guinardia, Dactyliosolen, Odontella and Lithodesmium) succession in Offatts Bayou, an embayment of Galveston Bay, Texas (Fig. 1). These eight centric planktonic diatom genera were chosen for their ease of identification and enumeration. They were also chosen because they dominated on the spatial (1 km^) and temporal (monthly) scales which are typically examined. Shifts in diatom community composition that are not related to natural variables (temperature, salinity) would appear useful as indicators of changing environmental conditions. Materials and Methods Study ^/Y^.-Offatts Bayou was created on Galveston Island early in the twentieth century after the 1900 Hurricane. The area was used as a borrow pit for local construction sites, in particular, the QUIGG & ROEHRBORN 285 seawall which now protects one-third of the seaward side of Galveston Island. By the end of the 1960's, Offatts Bayou was subject to extensive human development and became surrounded by homes and businesses, including Houston's most visited tourist attraction, Moody Gardens. Offatts Bayou is now approx. 1 km wide and 5 km long (Fig. 1). It is generally shallow (5-6 m, but mostly less), has poor circulation and restricted exchange with West Bay, its link to Galveston Bay (Fig. 1). Along its northern side is a boat channel which runs west to east; this side of the bayou is generally deeper (> 7 m). Most of the south side is not accessible by boat because of its shallow depths (< 1 m). Hence, sampling was performed along the northern bank with stations indicated on Fig. 1. Station 1 was located in West Bay at the mouth of Offatts Bayou; the remaining stations were spread approximately equidistant to the closed end finishing with station 6 (Fig. 1). The water column at West Bay (station 1) is well mixed in nature and relatively pristine (Roehrbom 2006). In the most restricted parts of the bayou (stations 5 and 6), residence times may be long (several months), water transparency is low (Secchi depths often 1-2 m less than those measured in West Bay (Skinner 2007), and the sediment and nutrient loading high as a result of dense development on its borders* Between these two extremes lies a gradient of water quality conditions and unique characteristics (bathymetry, mixing) that provides a range of environmental conditions for diatoms. Field procedures was performed over a 12 month period from December 2004 to November 2005. Phytoplankton tows (67 pm mesh net) were conducted at each station shown in Fig. 1 on a monthly basis. Water samples were transferred to 50 mL centrifuge tubes and stored in a cooler until returning to the laboratory. Phytoplankton were then preserved with formalin (final concentration 3%) and kept in a dark cool place until examined microscopically. Salinity and temperature were measured with a Hydrolab Mini Sonde (Hydrolab, Texas, USA) during field collections. All field sampling was conducted in the mornings between 0800 and 1 100 hrs Central Standard Time. 286 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 Table 1. Characteristics of the dominate diatom genera in Offatts Bayou between December 2004 to November 2005 at the six sampling stations. Species in this table reflect only those most commonly identified (> 90%) for each genera. As all diatoms were cylindrical in shape, biovolumes were calculated according to L*H*D*Pi/4. Genera Species {n > 20) Average length (nm) Average diameter (urn) Relative biovolume (pm^ ceirb Chaetoceros C. danicum 12-16 12-14 1,857 C. curvisetus 15-20 15-30 6,955 C. lorenzianus 30-48 12-24 9,920 Ditylum D. brightwellii 80-130 12-38 51,515 Rhizosolenia R. setigera 130-440 4-12 14,320 R. pungens 250-690 4-20 53,130 Coscinodiscus C. radiatus 25-35 45-55 58,875 Guinardia G. striata 50-80 5-12 3,690 G. delicatula 50 9-22 8,830 Dactyliosolen D. fragilissimus 8-70 42-300 964,076 Odontella O. mobilensis 35-90 24-70 108,379 O. sinensis 240-320 28-32 197,820 Lithodesmium L. undulatum 38-63 31-35 43,170 Phytoplankton analysis. — Light microscopy was used to identify and enumerate diatoms. Numerical abundance of diatom cells was determined by counting a minimum of five ocular micrometer grids at 400x magnification. The number of grids counted was adjusted according to cell density. Counts were completed upon reaching a minimum of 100 cells. The text used most frequently to aid in taxonomic identification was Tomas (1997). The primary focus of this study was the examination of centric planktonic diatoms which are reported only to the generic level. Because algal volumes can vary immensely among species, and because many ecological processes are more dependent on biovolumes than on densities, biovolumes were also estimated. Length and width measurements were calculated based on the averages (j7>20) determined for the species in Table 1. For example, all the Ditylum were D. brightwellii and the majority of Coscinodiscus were C. radiatus (>90%) so that biovolume calcula- QUIGG & ROEHRBORN 287 DJ FMAMJ J ASON DJ FMAMJ J ASON Month of the Year Figure 2. Monthly changes in surface water (a) temperature (°C) and (b) salinity (PSU) at the six stations in Offatts Bayou from December 2004 to November 2005. tions for these genera were based on dimensions of these species respectively (Table 1). Chaetoceros species were most often represented by C danicum, C. lorenzianus and C. curvisetus so that an 'average’ bio volume for Chaetoceros was calculated based on the mean measurements of these three species (Tables 1 & 2). It maybe for Chaetoceros that the calculated biovolume is underestimated given the sizes of these species. Biovolumes were calculated assuming all centric species were cylindrical: L*W*H*Pi/4 where L = length (pm), width (pm) and height (pm). Results Physical/chemical parameters ditox temperature did not vary spatially but did vary temporally (Fig. 2). Temperatures ranged from summer highs of 30.5°C (=tl°C) between June and August to winter lows of 14°C (±0.3°C) from December through the end of February. Salinity did not vary significantly among stations except for station 6 (ANOVA, >0.05), which had slightly lower salinities than the other stations. Salinities also varied temporally with highest salinities measured during July through to September (29 ± 0.3) and lowest salinities in the cooler months of December to March (19 ± 0.4) (Fig. 2). Table 2. Spatial distributions of the eight major diatom genera in Offatts Bayou between December 2004 to November 2005 at the six sampling stations. The biovolume of each genus at each station was calculated by multiplying the number of cells by the average biovolume for that genus. 288 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 I Q i a e) a ."5 "a I <3 s a Q C/D o OO (N o SO «n o r- in cn r-; (N o d d d cn m oo os r- Cs CN Os in CN| (N so’ in od OS so o O i> to p o VO d so iri d d SO os so VO m ' — ' ’—i o d d d d d so r- in o VO Os so d in (N ri o (N o^ 00 Os p so p o so os Os Os p r|- B 0 J3 (U o OC £ ’B O S cn 0 > p 6 0) > > .2 E H 3 zL < 15 zL QUIGG & ROEHRBORN 289 Month of the Year Figure 3. Spatial and temporal distributions of the eight dominate diatom genera in Offatts Bayou from December 2004 to November 2005. Diatom standing crops - abundance and biovolume. -Of the eight major planktonic diatom genera present in Offatts Bayou (Fig. 3), several different species of Chaetoceros accounted for 39% of the standing crop (numerically), while Ditylum accounted for 16%. Rhizosolenia (14%), Coscinodiscus (11%) and Guinardia (8%) also accounted numerically for relatively large fractions of standing crops while Dactyliosolen (5%), Odontella (4%) and Lithodesmium (4%) were present in the lowest proportions. Approximately 20 290 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 Figure 3. Cont. Other diatoms were present at the six stations during the sampling year; they accounted for < 1% of the total diatom community at anytime and were comprised of both centric and pennate forms, as well as other planktonic and some benthic types. Despite dominating numerically, Chaetoceros accounted for only 2% of the total biovolume of diatoms whilst Dactyliosolen accounted for the largest fraction (59%) (Table 2). Ditylum, Coscinodiscus and Odontella each accounted for -10% of the diatom biovolume in Offatts Bayou. QUIGG & ROEHRBORN 291 Diatom distributions - spatial patterns -DidXom standing crops of Chaetoceros, Ditylum, Odontella and Lithodesmium were highest at the mouth of Offatts Bayou, near West Bay, and then decreased along the sampling transect to the inner most station (Fig. 3) during the sampling period (December 2004 to November 2005). Wherever, Odontella and Dactyliosolen were present in high numbers, Guinardia and Lithodesmium were generally present in low numbers (Fig. 3). While this observation was not statistically significant, it was the pattern observed consistently over the study period as reflected in Fig. 3. Rhizosolenia and Coscinodiscus did not appear to exhibit any preferences spatially in Offatts Bayou. In the flow-restricted region of the bayou (stations 5 & 6), Guinardia and Dactyliosolen dominated. At station 1, where flow was not restricted but greatest mixing was generally observed, Odontella and Lithodesmium were present in highest quantities. Diatom distributions - temporal patterns diatom genera dominated at different times of the year (Fig. 3). While Chaetoceros dominated in the fall (September to November), a smaller peak was also observed late in the spring (March to June). These two peaks were associated with different species of Chaetoceros but clear species identifications were difficult with current resources. Ditylum and Rhizosolenia dominated late winter and early spring (December to March), while Coscinodiscus was prevalent between May and September (late spring to end of the summer). Each of these diatom genera appear to have dominated at most stations for several months or a season; during other times, they were still present in water samples, albeit at lower numbers. On the other hand, when Guinardia, Dactyliosolen, Odontella and Lithodesmium appeared, it was typically for about a month, usually at all stations, and then they essentially disappeared again from surface water samples (Fig. 3). Guinardia was present in highest densities in December, Dactyliosolen densities peaked from February to March, while Odontella and Lithodesmium were concurrently found in highest densities in from September to 292 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 November (Fig. 3). In this manner, different diatom genera peaked at different times of the year. Discussion Understanding natural changes in phytoplankton community composition is becoming more important with the increasing occurrence of eutrophication, changes in climate regulated processes which define circulation patterns, salinity regimes and pressures on ecosystems. Knowledge of the spatial and temporal distributions of estuarine phytoplankton offers insights into the potential effects of eutrophication and provides a tool for establishing relevant management strategies (Conley 2000; Olsen et al. 2001; Arhonditsis et al. 2007). Diatoms play an important role in many ecosystems (e.g., Badylak & Phlips 2004; Buyukates & Roelke 2005; Lavoie 2006) yet the bulk of studies using diatoms as bioindicators have been performed in freshwater systems. Most of the phytoplankton species identified in Offatts Bayou to date (Roehrbom 2006; Skinner 2007) were diatoms (up to 65 %). Similarly, Omolfsdottir et al. (2004a; 2004b) found diatoms dominated in Galveston Bay. They also found cyanobacteria, chrysophytes and cryptophytes were dominant at times while dinoflagellates, chlorophytes and euglenoids appeared occasionally abundant, but were relatively minor components on an annual basis. Buyukates & Roelke (2005) also found diatoms were an important component of the Nueces Delta (south Texas). While dinoflagellates and cyanobacteria are most often associated with eutrophication issues and are more likely to be associated with ‘harmful algal blooms’, the representatives in these taxa are often more difficult to identify and require more considerate handling and sampling methods. Because of the interest in developing a biomonitoring tool, diatoms were chosen for the reasons stated above and because of their established utility as biomonitors in other coastal systems (e.g., Resende et al. 2005; Cetinic et al. 2006) and in freshwater systems worldwide. QUIGG & ROEHRBORN 293 The eight dominant centric planktonic diatom genera examined in this study are cosmopolitan and can develop into bloom concentrations if conditions are suitable. Blooms of Dactyliosolen, Odontella, Chaetoceros, and Rhizosolenia for example, were observed in the Indian River Lagoon, Florida, USA (Badylak & Phlips 2004). Wang et al. (2006) found that Rhizosolenia and Chaetoceros dominated in Daya Bay (China). Although present, Skeletonema and Nitzschia were not observed in high abundances in Offatts Bayou (Roehrbom 2006); these genera are however known to dominate in other systems (Badylak & Phlips 2004; Buyukates & Roelke 2005; Wang et al. 2006). The eight planktonic genera examined in this study are not known to be harmM based on their absence from the 'harmful algal bloom’ list developed by the Intergovernmental Oceanographic Commission of UNESCO (IOC 2007). Spatial and temporal patterns in the diatom community - Similarly to earlier studies on Offatts Bayou (e.g., Cooper & Morse 1996), seasonal variations were observed in the thermohaline structure of Offatts Bayou over the course of the study (December 2004 to November 2005). Temperatures and salinities were also typical of those measured in subtropical bays and bayous (Quigg et al. 2007; Quigg et al. 2009) and lagoons (Badylak & Phlips 2004) in the Gulf of Mexico. Winter waters were cooler, less saline and mixed while summer waters were warmer, saltier and often stratified (Fig. 2; Roehrbom 2006). In Fig. 2, the importance of salinity and temperature in regulating the composition of this diatom community on temporal scales in Offatts Bayou is seen. It is well know that other factors such as light, nutrients, grazing, mixing, would also be important. These however, were not measured comprehensively as part of this initial investigation due to insufficient resources. Light, nutrients and mixing will be measured in omgoing studies. Adaptation to osmotic stress is a major factor in dictating the success of a particular species in estuarine environments, and can have broad impacts on phytoplankton composition and standing crops (Kennish 2002). 294 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 Resende et al. (2005) and Cetinic et al. (2006) recently also reported that salinity and temperature were the most important factors in defining species composition in the Canal de Mira, Ria de Aveiro (Western Portugal) and in the Krka Estuary (Adriatic coast), respectively. Hence, these two measures alone may not describe in full detail the pressures on the community, but are nonetheless important. Three possible scenarios may explain the relatively low diatom standing stocks in summer in Offatts Bayou compared to the rest of the year. First, small rapidly growing diatoms may be replaced by slow-growing, filamentous or inedible forms of diatoms and other phytoplankton during low-nutrient conditions such as those that occur in summer as a result of little or no rainfall, and/or freshwater inputs (Sommer 1989). Although, nutrients were not measured as part of this study, earlier findings revealed that nitrate concentrations were below detection (year round), phosphate varying from zero (winter) to 3 mM (summer) and silicate from zero (winter) to 30 uM (summer) in surface waters (top 2-3 m) of Offatts Bayou (Cooper & Morse 1996). Similar nutrient distributions have been observed for Galveston Bay (Santschi 1995; Quigg et al. 2007). While the actual concentrations during this study may not have been the same, similar patterns were likely, particularly the low nutrient concentrations in the summer. Winter diatom populations were likely to be N-limited given the phosphate and silicate concentrations. This is in fact what was observed. Studies of the constraints on primary productivity in Galveston Bay also found the system to be predominately N-limited (Omolfsdottir et al. 2004; Quigg et al. 2007). Changes in relative abundances of diatoms under increasing nitrogen loading and/or other human- induced changes to estuarine hydrography thus have important implications for Offatts Bayou and other estuaries. Second, centric diatoms are nonmotile and prone to sinking out of the water column (Smetacek 1999). Sinking may be exacerbated in the summer in Offatts Bayou given its highly stratified nature QUIGG & ROEHRBORN 295 during the warmer months (Roehrbom 2006; Skinner 2007), much like that which occurs in other subtropical estuarine systems (Pinckney et al. 2001; Paerl et al. 2003; Wang 2006). Hurricanes, such as Rita, which came close to Offatts Bayou in the summer of 2005 (September 24), may temporarily inject the system with nutrients and/or disrupt stratification. It was noted however, that stratification had been re-established within two weeks of Hurricane Rita passing. This was surprising given that recorded wind gusts at the nearby meteorological station (Scholes airport; Fig. 1) were >100 km h"^ for many hours. Third, the timing of the decrease in diatom biomass during summer may have coincided with the timing of greatest zooplankton grazer populations in this system. Although zooplankton samples was not quantified during this study, larger numbers of grazers were observed to be present. Also, phaeophytin concentrations, which are sometimes used as a proxy for grazing pressure, were elevated in the summer months (Roehrbom 2006; Skinner 2007). Large centric planktonic diatoms such as those investigated in the present study are an important part of the diet of many copepod species, which are in turn preyed upon by fish, including important commercial fish species. Thus any change in the abundance of centric diatoms could cause changes in the food web stmcture in this and/or other estuarine systems. Whether results were presented as relative abundance (Fig. 3) or biovolume (Table 2), the overall findings in terms of spatio- temporal distributions of these planktonic diatoms were the same. Given the inherent difficulties associated with calculating cell size (biovolume and surface area), the additional total analysis time (substantial) due to the numerous measurements required, and the possibility of overestimating the contribution of a particular genera or species with large vacuoles to the overall community, the findings of this study are presented predominantly on the basis of diatom abundances (Fig. 3). Lavoie et al. (2006) came to a similar 296 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 conclusion, determining that relative abundance was the most appropriate metric to use for biomonitoring purposes. This study provides a snapshot of the diatom community structure in Offatts Bayou, Texas. This is a valuable dataset in providing a baseline on the nature of this system, especially given that eutrophication, development and climate change will concurrently modify it in the future. Ongoing studies are being conducted in Offatts Bayou and Galveston Bay with the intent to develop microbial indicators for these aquatic ecosystems. Future studies will involve collecting nutrient data and ultimately modeling efforts such as those presented in Arhonditis et al. (2007). Such tools are required for the development of effective eutrophication management strategies. Acknowledgements We thank all the undergraduate and graduate students that participated in the field work, in particular, Allison Skinner, Derek Shultz and Federico Alvarez. Amber Bradford helped with phytoplankton identification and enumeration. Daniel Roelke and Jay Pinckney are thanked for comments and suggestions. The junior author acknowledges support throughout her Masters program from the Texas Institute of Oceanography and Texas A&M University at Galveston. We thank reviewers Roy Lehman, Hudson Deyoe and the Manuscript Editor for their comments. Literature Cited Arhonditsis, G. B., C. A. Stow, H. W. Paerl, L.M. Valdes- Weaver, L. J. Steinberg & K. H. Reckhow. 2007. Delineation of the role of nutrient dynamics and hydrologic forcing on phytoplankton patterns along a freshwater-marine continuum. Ecol. Modeling, 208:230-246. Badylak, S. & E. J. Phlips. 2004. Spatial and temporal patterns of phytoplankton composition in a subtropical coastal lagoon, the Indian River Lagoon, Florida, USA. J. Plank. Res., 26:1229-1247. Buykates, Y. & D. L. Roelke. 2005. Investigating system characteristics of a southwest Texas wetland: Nutrient and plankton dynamics of a tidal creek in the lower Nueces delta. J. Appl. Sci., 5:820-828. Cetinic, L, D. Vilicic, Z. Buric, & G. Olujic. 2006. Phytoplankton seasonality in a highly stratified karstic estuary (Krka, Adriatic Sea). HydrobioL, 555:31-40. QUIGG & ROEHRBORN 297 Conley, D. J. 2000. Biogeochemical nutrient cycles and nutrient management strategies. Hydrobiol., 410:87-96. Cooper, C. D. & J. W. Morse. 1996. The chemistry of Offatfs Bayou, Texas: a seasonally high sulfidic basin. Estuaries, 19:595-61 1. Fryxwell, G. A,, M. E. Reap & D. L. Valencic. 1990. Nitzchia pungens Grunow F. miiltiseries Hasle: observations of a known neurotoxin diatom. Nova Hedwigia., 100:171-188. Galveston Bay Estuary Program (GBEP) 2001. The Galveston Bay Plan. The comprehensive conservation and management plan for the Galveston Bay Ecosystem. GBNEP#49. 457 pp, Hallegraeff, G. M. 1993. A review of harmful algal blooms and their apparent global increase. Phycologia, 32:79-99. Houston-Galveston Area Council (H-GAC). 2006. Basin Summary Report-2006. Houston, Texas. Howarth, R., D. Anderson, J. Cloem, C. Elffing, C. Hopkinson, B. Lapointe, T. Malone, N. Marcus, K. McGlathery, A. Sharpley & D. Walker. 2000. Nutrient Pollution of Coastal Rivers, Bays, and Seas. Iss. in EcoL, 7:1-17. IOC 2007. IOC taxonomic reference list of toxic algae. Intergovernmental Oceanographic Commission of UNESCO. Available from: www.ioc.unesco.org/hab, last accessed October 12, 2007. Kennish, M. J. 2002. Environmental threats and environmental future of estuaries. Environ. Conserv., 29:78-107. Lavoie, I., S. Campeau, M-A. Fallu & P. J. Dillon. 2006. Diatoms and biomonitoring: should size be accounted for? Hydrobiol., 57:1-16. Lester, L. J. & L. Gonzalez (Eds.). 2002. The State of the Bay: A Characterization of the Galveston Bay Ecosystem. GBEP T-7. Webster, Texas. Nixon, S. W. 1995. Coastal marine eutrophication: a definition, social causes and future concerns. Ophelia, 41:1 99-2 1 9. Olsen, Y., H. Reinertsen, O. Vadstein, T. Andersen, 1. Gismervik, C. Duarte, S. Agusti, H. Stibor, U. Sommer, R. Lignell, T. Tamminen, C. Lancelot, V. Rousseau, E. Hoell & K. A. Sanderud. 2001. Comparative analysis of food webs based on flow networks: effects of nutrient supply on structure and function of coastal plankton communities. Cont. Shelf Res., 21:2043-2053. Omolfsdottir, E. B., S. E. Lumsden & J. L. Pinckney. 2004a. Nutrient pulsing as a regulator of phytoplankton abundance and community composition in Galveston Bay, Texas. J. Exp. Mar. Biol. EcoL, 303:197-220. Omolfsdottir, E. B., S. E. Lumsden & J. L. Pinckney. 2004b. Phytoplankton community growth-rate response to nutrient pulses in a shallow turbid estuary, Galveston Bay, Texas. J. Plankt. Res., 26:325-39. Paerl, H. W., J. L. Pinckney, J. M. Fear & B. J. Peierls. 1998. Ecosystem responses to internal and watershed organic matter loading: consequences for hypoxia in the eutrophying Neuse River Estuary, North Carolina, USA. Mar. Ecol. Prog. Ser., 166:17-25. Paerl, H. W., Valdes, L. M,, Pinckney, J. L., Piehler, M. F., Dyble, J. & Moisander, P. H. 2003. Phytoplankton photopigments as indicators of estuarine and coastal eutrophication. Biosci., 53:953-64. 298 THE TEXAS JOURNAL OF SCIENCE, VOL. 60, NO. 4, 2008 Pinckney, J. L., H. W. Paerl, P. A. Tester & T. L. Richardson. 2001. The role of nutrient loading and eutrophication in estuarine ecology. Environ. Health Pers., 109:699-705. Quigg, A., S. E. Davis & D. F. Roelke. 2007. Changes in Freshwater Inflows and How They Effect Texas Bays. Final Report CCC, NOAA No. NA05NOS- 4191064, 47 pp. Quigg, A., L. Broach, W. Denton & R. Miranda. 2009. Water quality in the Dickinson Bayou Watershed (Texas, Gulf of Mexico) and health issues. Marine Pollution Bulletin (in press). Access at: http://dx.d0i.0rg/l 0. 1 0 1 6/j .marpolbul.2009.0 1.012 Reap, M. E. 1991. Nitzchia pungens Grunow F. multiseries Hasle: Growth phases and toxicity of clonal cultures isolated from Galveston, Texas. Masters Thesis. Texas A&M Univ., College Station, 77 pages. Resende, P., U. Azeiteiro & M. Jorge Pereira. 2005. Diatom ecological preferences in a shallow temperate estuary (Ria de Aveiro, Western Portugal). Hydrobiologia, 544:77-88. Roehrbom, L. 2006. Seasonal analysis of abiotic factors impacting phytoplankton assemblages in Offatts Bayou, Galveston, Texas. Unpublished MS Thesis. Texas A&M Univ., College Station, 273 pp. Santschi, P. H. 1995. Seasonality in Nutrient Concentrations in Galveston Bay. Mar. Environ. Res., 40:337-62. Skinner, A. C. 2007. Belly-up in the Bayou, who’s the culprit? Physical, Chemical, and Biological parameters of Offatts Bayou, Galveston, Texas. Unpublished Honors Thesis. Texas A&M Univ., Galveston, 64 pp. Smetacek, V. 1999. Diatoms and the ocean carbon cycle. Protist, 150:25-32. Sommer, U. 1989. The role of competition for resources in phytoplankton succession. Pp. 57-106, in Sommer, U. [Ed.] Plankton Ecology: Succession in Plankton Communities. Springer- Verlag, Berlin, 369 pp. Texas Water Development Board. 2007. State Water Plan for Texas: Vol. I. Highlights of the 2007 State Water Plan. Document no. GP-8-1. Thronson, A. & A. Quigg. 2008. Fifty-five years of fish kills in Coastal Texas. Estuaries and Coasts, Vol. No. 31:802-813. Tomas, C. R. 1997. Identifying Marine Phytoplankton. Academic Press, California, 858 pp. Wang, Y-s., Z. P. Lou, C. C. Sun, W. L. Wu & S. H. Han. 2006. Multivariate statistical analysis of water quality and phytoplankton characteristics in Daya Bay, China, from 1999 to 2002. Oceanologia, 48:193-21 1. AQ at: quigga@tamug.edu TEXAS J. OF SCI. 60(4):299-308 NOVEMBER, 2008 AN EVALUATION OF MODIFIED IMS SWABS FOR THE SCREENING OF OXIDIZERS AND HOME-MADE EXPLOSIVES Karl D. Kuklenz and Darren L. Williams Chemistry Department, Sam Houston State University Huntsville, Texas 77340 Abstract-Two cotton wipe treatments were developed, and a commercial colorimetric wipe was evaluated with the goal of improving airline security screening of suspect articles for chemical oxygen generators (COG) and homemade peroxide- containing explosives (HME). Five redox indicators (2,3’-diphenylamine dicar- boxylic acid, diphenylaminesulfonic acid, diphenylamine, iodine: starch, and luminol) were evaluated against hydrogen peroxide and potassium chlorate. Diphenylamine and iodine: starch worked in the dry state but with very slow color-development times - negating their effectiveness as a primary security screening technique. The use of a color-developing spray reduced the detection times of the formulations prepared in this study. A commercial product (Drop-Ex Plus, Mistral Security Inc.) contained color developing solvents and produced similar results. The need for color- developing solvents relegates the colorimetric wipe systems to secondary screening where separate wipes can be used for the nitro-based explosives, COG, and HME. With the constant threat of terror attacks present in the minds of world travelers it is important to have a fast, cost-effective, and safe way to individually screen luggage and carry-on items for explosive residues and hazardous substances. The current primary security screening method employs ion mobility spectrometry (IMS) to detect nitro-containing explosives such as trinitrotoluene (TNT) and pentaerythritol tetranitrate (PETN) (Ewing et al. 2001) in approxi¬ mately six seconds per screened article (DHS S&T 2006). Liquids in containers with a capacity greater than 59 mL (two fluid ounces) are currently confiscated because the TSA has not developed or deployed a primary screening procedure for chemical oxygen generators (COG) and peroxide-containing explosives (FAA 2007). The typical single-tube IMS is set to detect negative ions because the volatile nitro-containing explosives readily form negatively-charged adducts with the reactant gas in the IMS ionization chamber, as illustrated: H-R-NO2 (g) + Cf (g) [Cl (H- R-NO2)]" (g). The chloride-laden reactant gas is produced by beta- 300 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 particle-induced dissociation of methylene chloride inside the IMS. (Eiceman & Stone 2004) Triacetonetriperoxide (TATP) and hexamethylene triperoxide diamine (HMTD) are homemade explosives (HME) that are becoming more widely used in attacks (Buttigieg et al. 2003), These peroxide-based explosives are dangerous to synthesize, but large amounts can be made from inexpensive items such as acetone, sulfuric acid, 3% hydrogen peroxide, citric acid, and hexamethy¬ lenetetramine (Shakashiri 1983). The triperoxides form positive-ion adducts in the IMS and are not detected by single-tube IMS instruments in negative-ion mode (Buttigieg et al. 2003). Dual-mode instruments are now available that can detect HMEs and traditional explosives simultaneously. However, more than 10,000 single-mode IMS instruments are currently deployed at airports worldwide (Eiceman & Karpas 2005), and the expense of replacement with dual-tube IMS as well as commensurate training of IMS operators would significantly postpone large-scale implementation. Even so, current IMS will only detect volatile nitro-based explosives. An inexpensive screening technique that detects oxidizers (i.e. peroxides and chlorates) is needed (FAA 2001). Since most airport security screening is based upon a wipe sampling technique (Eiceman & Stone 2004), a dry peroxide¬ detecting colorimetric wipe that does not interfere with the single¬ tube IMS screening procedure would be a valuable interim solution. Ideally in primary screening, the security official would swab the suspect article and look for a color change on the swab. If the color indication were negative, the official would use the same swab in the IMS to detect nitro-containing explosives. If the color indication were positive, the article would then be pulled aside for further inspection. A dry swab is also desired as this would minimize interference with the IMS detection of nitro explosives. KUKLENZ & WILLIAMS 301 In a secondary screening scenario, normal TSA procedures would identify suspect articles to be wiped with the colorimetric swab as a test for the presence of HME or COG residues. Time constraints for secondary screening are not as restrictive as those for primary screening, so a color-development time could be tolerated. Also, IMS interference is not an issue since a different wipe could be used for the primary nitro-based explosive screening. Concentrated peroxides and COG have been banned from passenger air transport since the 1996 ValuJet accident (FAA 2007; 49 CFR §172.101). A good screening technique for oxidizers will also detect peroxidic HMEs. The TSA has expressed interest in better screening and identification methods for oxidizers (FAA 1998; 2001) that are banned from commercial aircraft (Table 1). The CRC Handbook (Tide & Frederikse 1996) lists the standard reduction potentials of these substances. A high reduction potential represents a strong oxidizing agent. Several colorimetric redox indicators are given in Table 2 along with the color of the oxidized species. Skoog et al. (1996) report that redox indicators typically depend only upon the cell potential of the system, which is the reduction potential of the oxidizing species minus the reduction potential of the indicator. This reaction is spontaneous when the standard reduction potential of the oxidant is greater than the standard reduction potential of the indicator. It is anticipated that the standard cell potentials will accurately reflect the spontaneity of the reactions in the solid state. Many of the indicator reactions require water, hydroxide ions, or acid protons as reactants (Skoog et al. 1996), but the spontaneity of these reactions should allow the oxidizing agents to utilize the water, protons, and hydroxyl groups present in the cotton fiber matrix. If the rate of mixing of the reactants in a dry wipe is slow, it may be necessary to employ a liquid developing solution. 302 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Table 1. TSA forbidden oxidants and explosives. Substance Bred (V) Triacetone triperoxide (TATP) explosive Hexamethylene triperoxide diamine (HMTD) explosive Hydrogen peroxide, H2O2 +1.8 Chlorate, CIOs' +1.5 Permanganate, Mn04' +1.5 Perchlorate, C104' +1.4 Dichromate, Cr207^' +1.2 Hypochlorite, CIO' (Bleach) +0.8 The same cotton swabs used in the lonscan 400B IMS (Smiths Detection, Inc) were treated with the redox indicators in Table 2 and were evaluated both dry and wet to determine if colorimetric wipes were suitable for primary or secondary security screening. Hydrogen peroxide was tested in this study as a representative of the peroxidic explosives, and potassium chlorate was tested as a representative of the chemical oxygen generators. Materials and Methods The indicators 2,3’-diphenylamine dicarboxylic acid, diphenylamine sulfonic acid, and diphenylamine were exposed to potassium chlorate to investigate the result as it applies to visual detection and screening. The individual diphenylamine indicators were prepared as methanol solutions with 1% w/w indicator. One milliliter of the 2,3 ’-diphenylamine dicarboxylic acid indicator was pipetted into each of three 1.5 mL vials with caps. This was repeated for the other two indicator solutions for a total of nine vials. Each set of three indicators was probed with 100 pL of three different concentrations of potassium chlorate (0.1 M, 0.05 M, and 0.01 M). The solutions were then developed with two drops of hydrochloric acid. The results were evaluated subjectively on the basis of opacity, color, and development time. Based on these KUKLENZ & WILLIAMS 303 Table 2. Redox indicators tested in this study. Substance Indicating color Bred (V) 2,3’-diphenylamine dicarboxylic acid diphenylamine sulfonic acid diphenylamine iodine: starch luminol chemiluminescent Blue-black blue-violet red-violet violet +1.12 +0.85 +0.76 +0.54 results the indicator diphenylamine was selected as a candidate for dry wipe testing. Three indicator solutions were prepared for dry wipe testing. Indicator solution #1 was prepared as a methanol solution with 1% w/w diphenylamine and 5% hydrochloric acid. Indicator solution #2 was prepared as a 0.1 M potassium iodide aqueous solution containing 1% w/w starch. Indicator solution #3 was a luminol indicator prepared from the following reagents: 0.1% sodium carbonate, 1% sodium bicarbonate, 0.01% ammonium carbonate monohydrate, 0.01% copper (II) sulfate, and 66 ppm of 3- aminophthalhydrazide (luminol). The recipe for indicator #3 was taken from Shakashiri (1983). Three IMS swabs (Smiths Detection Inc.;one for chlorate testing, one for peroxide testing, and one for a control) were soaked in each of the indicator solutions. The control would be evaluated for resistance to atmospheric oxidation. The swabs were allowed to dry overnight at room temperature and atmospheric pressure. A surface was prepared on a clean epoxy resin countertop. An area of 7 by 10 cm was masked off for each swab. A 50 pL drop of either 0.01 M potassium chlorate or 3% hydrogen peroxide was placed in the center of each area and allowed to dry for 5 h. This yielded a contamination of approximately 61 pg KCIO3 and 2.2 pg H2O2 upon drying. 304 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 When both the modified swabs and the test areas were dry a separate swab was used for each test area. The full test area was wiped, and the color change was evaluated over time. The experiment was repeated in its entirety to evaluate the effect of a moistening step after dry-wiping the test area. The modified swabs were moistened with a single spray of water from a common pneumatic pump spray bottle. A commercial off-the-shelf (COTS) detection package - Drop- Ex Plus (Mistral Security Inc.) - was tested for the detection of hydrogen peroxide and potassium chlorate residues. The kit provided collection paper and analysis reagents in drop-dispensing bottles. The kit also came with a procedure and chart for analyzing the results. A surface was prepared on a clean countertop as before. Test areas were masked off and similar peroxide and chlorate residues were applied. After the surface reagents were dry a collection paper was taken from the Drop-Ex Plus kit and used to swab one of the test areas. Two drops of the proprietary Drop-Ex Plus chlorates indicator (bottle A) were applied to the test paper to test for chlorate residues. If a dark-blue color change was not observed, the test was continued for peroxide residues on the same wipe. Two drops of the proprietary Drop-Ex Plus peroxides indicator (bottle B) were applied to the collection paper. If a dark- blue color change occurred at this point the presence of peroxides was suggested. This experiment was repeated twice for a total of three experiments. The Drop-Ex Plus kit verification sample was tested, thus ensuring that the indicator solutions were behaving as expected. Results and Discussion The diphenylamine species comparison tests were subjectively graded on opacity, color, and development time. The 2,3’- diphenylamine dicarboxylic acid reacted in solution to produce a translucent red-brown color 15 sec after addition of the acid developer. The intensity of the red-brown color increased with KUKLENZ & WILLIAMS 305 Table 3. COG and HME detection results of the treated wipes Indicator Analyte Solution Response Time (min) Dry Response Time (min) Wet Response Time (min) 2.3 ’-diphenylamine chlorate 0.25 - - dicarboxylic acid peroxide - - - diphenylamine chlorate 0.25 - - sulfonic acid peroxide - - - diphenylamine chlorate instantly 15 3 peroxide - - - Kl/starch chlorate - - - peroxide instantly 35 5 luminol chlorate - - - peroxide instantly - - Drop-Ex Plus chlorate - - < 1 peroxide - - < 1 increasing chlorate ion concentration. Diphenylamine sulfonic acid performed in a similar fashion, leaving a light, translucent green solution 10 - 15 sec after development. In contrast, the indicator diphenylamine developed immediately after the addition of hydrochloric acid. All three vials were an intense navy-blue. This color was intense enough to prevent light from passing through the side of the small vial. Because of its fast development time and unmistakable color, diphenylamine was chosen for dry-wipe testing. Dry, treated wipes modified with the luminol indicator (#3) showed no chemiluminescence in ambient room light or in total darkness (Table 3). Swabs treated with iodine:starch (#2) were able to detect peroxides but only after a swab development time of 30 min or more (Fig. 1). These swabs were totally ineffective at screening chlorate residues from the surface no matter how long they were allowed to develop. Swabs treated with diphenylamine (#1) were able to detect chlorate residues but only with 306 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Figure 1. A representative result of the iodine: starch indicator (#2) detecting the presence of peroxides (left) and the untreated control wipe (right). development times of 15 min or more. These swabs were totally ineffective at detecting peroxide residues, even after a long development time. In summary, the rate of mixing of the dry reactants was too slow to detect COG and HME residues in primary screening. The addition of water spray to the procedure significantly reduced development times to 5 min for indicator #2 detecting peroxide residues and to 3 min for indicator #1 detecting chlorate residues. Indicator #2 still was not able to detect chlorate residues and indicator #1 still was not able to detect peroxide residues. Indicator #3, the luminol reagent was not able to detect either species in ambient room light or even when modified swabs were inspected in a dark room. The Drop-Ex Plus kit was able to successfully screen all chlorate and peroxide residues on the surface. Sample collection, testing, and analysis were performed in less than 1 min, and the results were clear and easy to evaluate. Because the collection paper is not KUKLENZ & WILLIAMS 307 chemically treated and reagents are stored as liquids in plastic bottles, atmospheric oxygen interference was not an issue. Conclusions The wipe and look procedure achieved by a chemically-modified dry-wipe will not work for primary security screening because of the limited reactant mobility and long color-development times. The use of a developing solution shortens this time, but excess moisture would not be compatible with the subsequent IMS detection of the nitro-based explosives. Therefore, the wipe treatments and developing solutions described herein are useful only for secondary screening for COG and HME. In summary, the wipe-treatment preparation instructions in this article would allow production of COG- and HME-detection wipes for secondary screening at low cost. Potassium chlorate and 3% hydrogen peroxide solutions would be suitable for use as verification solutions. A COTS solution similar to the Drop-Ex Plus detection kit would also work well for detection of oxidizers and explosives. This kit contains the required test procedures and verification samples, and it performed as well as the wipes prepared in this study. Acknowledgements The Welch Foundation Department Development Grant and the Sam Houston State University Enhancement Grant for Research programs are acknowledged for their financial support of this work. Laurie Raesz is also acknowledged for her assistance preparing the treated wipes. Literature Cited Buttigieg, G, A., A. K. Knight, S. Denson, C. Pommier & M. B. Denton. 2003. Characterization of the Explosive Triacetone Triperoxide and Detection by Ion Mobility Spectrometry, Forensic Sci. Int., 135: 53-59. Code of Federal Regulations, Title 49: Transportation, Section 172.101 Hazardous Materials Table, http://eCFR.GPOAccess.gov/ (accessed June 2008) 308 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Department of Homeland Security Science and Technology Directorate (DHS S&T), Broad Agency Announcement BAA-06-00063 Proposal Information Pamphlet, 2006, §3.1, 6-7. Eiceman, G. A. & Z. Karpas. 2005. Ion Mobility Spectrometry 2"^ Ed. CRC Press, Boca Raton, 350 pp. Eiceman, G. A. & J. A. Stone. 2004. Ion Mobility Spectrometers in National Defense, Anal. Chem. A Pages, 390A-397A. Ewing, R. G., D. A. Atkinson, G. A. Eiceman, G. J. Ewing. 2001. A Critical Review of Ion Mobility Spectrometry for the Detection of Explosives and Explosive Related Compounds, Talanta, 54: 515-529. Federal Aviation Administration, Civil Aviation Security Dangerous Goods Advisory Bulletin, Prohibition of Oxygen Generators (Chemical) Aboard Passenger Aircraft, DGAB-96-01, 1998, 1-2. Federal Aviation Administration, Civil Aviation Security Dangerous Goods Advisory Bulletin, Transportation of Hydrogen Peroxide (H2O2), DGAB-01-01, 2001, 1-2. Federal Aviation Administration (FAA), Civil Aviation Security Dangerous Goods Advisory Bulletin, Chemical Oxygen Generators (COGs) and Chemical Oxygen Generators Installed in Equipment, DGAB-07-02, 2007, 1-3. Lide, D. R. & H. P. R. Frederikse, Eds. 1996. CRC Handbook of Chemistry and Physics, 76th ed.; CRC Press, New York, NY, §8, 21-33. Shakhashiri, B. Z. 1983. Chemical Demonstrations: A Handbook for Teachers of Chemistry. Vol. 1. The University of Wisconsin Press, pp. 46-48. Skoog, D. A., D. M. West & F. J. Holler. 1996. Funtamentals of Analytical Chemistry 7th Ed.; Saunders College Publishing, New York, NY, 354-358. DLW at: Williams@shsu.edu TEXAS J. SCI. 60(4), NOVEMBER, 2008 309 GENERAL NOTES NOTES ON BEHAVIOR OF THE TEXAS KANGAROO RAT {DIPODOMYS ELATOR) Jim R. Goetze*, Allan D. Nelson and Chad Stasey ^Science Department, Laredo Community College Laredo, Texas 48040 and Department of Biological Sciences, Tarleton State University Box T~0100, Stephenville, Texas 76402 The Texas kangaroo rat {Dipodomys elator) is listed as a threatened species by the Texas Parks and Wildlife Department (Schmidly 2004). Reasons for listing D. elator are largely based on its apparent scarcity and small geographic range. Investigators have found D. elator in only 10 counties in north-central Texas (Schmidly 2004). The International Union for Conservation of Nature (1986) listed habitat loss and degradation resulting from expanding agricultural and infrastructure development as the major threats to continued existence of D. elator. There is general agreement that D. elator requires a sparse, short-grassland habitat (Carter et al. 1985; Dalquest & Collier 1964; Goetze et al. 2007; Roberts & Packard 1973; Stangl et al. 1992), but this type of habitat is becoming less common throughout the present range of the Texas kangaroo rat (Goetze et al. 2007). With the exception of a few brief reports (Dalquest & Collier 1964; Chapman 1972; Roberts & Packard 1973; Packard & Roberts 1973), little information exists on behavior of the Texas kangaroo rat. Information on home ranges and burrow affinity was mostly obtained in catch-and-release studies of this Texas state-threatened species. Sociality and other behaviors have been described for other species of kangaroo rats (Behrends et al. 1986; Randall 1989; 1991; 1993; 2007; Jones 1993; Yoerg 1999), but have not been specifically documented for D. elator. A thorough understanding of the social interactions and other behaviors of a species provides vital information for its conservation (Allison & Destefano 2006). 310 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Focal observations on agonistic, dust bathing, foraging, and day burrow usage behaviors for D. elator are presented within this note. During a study involving habitat characterization, burrow use, and population estimation of the Texas kangaroo rat (Goetze et al. 2007), night vision scopes were used to closely observe behavior. Dipodomys elator is a noctumally active species and previous researchers have utilized dim incandescent lighting to observe behavior. The use of infrared scopes disturbs the animals less and is an additional way of observing behavior of this species. In their study of equipment and techniques for nocturnal wildlife studies, Allison & Destefano (2006) pointed out that night vision scope generation model, light pollution due to nearness of urban areas, approach distance to study subjects, ability to distinguish individuals, and inclement weather were potential drawbacks of research utilizing night vision scopes. However, the D. elator study site was not located near an urban area, therefore light pollution levels were low. Also, researchers were able to approach to within a few meters of Texas kangaroo rats without significant disturbance, therefore individuals could easily be focally observed. Only two observation periods were interrupted by inclement weather. Because kangaroo rats habituate readily to observers (Randall 1991) and D. elator usually occurs in areas with little concealing vegetation, the Texas kangaroo rat proved to be a good subject for study utilizing night vision scopes. The 15 ha study site is located in Wichita County, Texas, 3 km N Buffalo Creek Reservoir and was previously described by Goetze et al. (2007). The study area is heavily grazed and bordered by wheat fields on its eastern and western sides, and contains short- grasses and low shrubs. Mechanical brush control and herbicide applications have resulted in small (1-2 m in height) and scattered woody vegetation. These habitat features facilitated observations with the night scopes. TEXAS J. SCI. 60(4), NOVEMBER, 2008 311 Observations were made utilizing a pair of Night Owl Optics model NOLT3 Generation One monocular night scopes. Fourteen observational sessions were conducted during the months of May and June, 2004 and 2005, for a total of 101 hours and 15 minutes. Observation began either before sunset or, in two cases, during early morning hours (330 and 400 hrs.). One observational period extended from sunset to sunrise. Behavioral observations were recorded in field journals. Focal observations were conducted at a distance of approximately three to five meters from the animals. However, observations of agonistic behaviors and dust bathing were often made at closer distances. Agonistic behavior -Agonisiic behavior of D. elator has only been reported from a laboratory setting between males and females (Packard & Roberts 1973). Females harassed males by nipping them on the bulky portion of the body. In this study, agonistic behavior was observed in the field when a striped skunk {Mephitis mephitis) ran down a fence-row trail near two foraging kangaroo rats causing the rats to come together. These kangaroo rats jumped into each other several times and pushed away in mid-air until one antagonist disengaged and hopped back toward a burrow. Later, one of the previous individuals hopped to another individual's burrow as the preceding individual foraged some distance away. The intruder dug up seed caches at the burrow entrance and around the base of a shrub until the burrow’s occupant returned and aggressively contacted the intruder and chased this rat down an established trail toward its own burrow. These contacts occurred three times during the observational period. Pilfering of seed caches has also been observed in Merriam's kangaroo rat {D. merriami) (Daly et al. 1992). An additional incidence of agonistic behavior was observed between two individuals who fought at a dust bathing site along a trail connecting their burrows. The conflict occurred at a point approximately midway (14 m) between the two burrow entrances. In each instance, the intruding individual retreated back to its own burrow location. Texas kangaroo rats may forage in distinct, defended territories along their trails. When one 312 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 kangaroo rat entered another’s territory, the resident leapt into the air several times possibly indicating to the intruder that conflict was eminent. If the intruder did not retreat, the resident sometimes engaged the intruder and both individuals bumped and pushed against each other. If the conflict continued, the antagonists rolled and scuffled on the ground until one retreated. Similar agonistic behavior has been reported for D. heermanni and D. merriami (Randall 1989; Yoerg 1999). Texas kangaroo rats occasionally forage in areas away from established trails and dust bathing sites within two to five meters of each other without conflict. Such behavior has been observed in Merriam's kangaroo rat (Behrends et al. 1986). Dalquest & Collier (1964) and Packard & Roberts (1973) never observed two or more adult D. elator above ground together. Observations made during this study may indicate that agonistic behavior occurs only within well-defined territories such as dust bathing sites and trails associated with burrows. Behrends et al. (1986) and Jones (1993) have also reported territorial behaviors in D. merriami, D. heermanni, and D. spectabilis. Dust bathing-\]^Q of scratching and dusting areas by D. elator has been reported by other researchers (Dalquest & Collier 1964; Packard & Roberts 1973; Carter et al. 1985), but none have described dust bathing behavior for the species. Several D. elator conducting dust bathing activities were closely observed. Dust bathing occurred in bare areas containing loose, fine-grained soils immediately adjacent to trails. Dust bathing was observed in five individuals. The kangaroo rats scratched in the soil with their hind legs, rubbed their sides and their bellies in the dust and stroked the sides and front of their faces at and around the cheek pouches with the front feet. This behavior is similar to dust bathing activities reported for other species of heteromyids by Randall (1991; 1993). Deposition of oils from the dorsal skin gland at dust bathing sites plays a role in neighbor recognition, at least in some species of kangaroo rats (Randall 1991; 1993). Although this aspect of TEXAS J. SCI. 60(4), NOVEMBER, 2008 313 behavior in D. elator was not observed during this study, Stangl et al. (2006) found that size and development of the dorsal gland of the Texas kangaroo rat was correlated with an individual's reproductive condition. Foraging behavior behaviors were observed 12 times. Foraging kangaroo rats moved forward slowly upon all fours and utilized their forelimbs to place food in their cheek pouches, instead of immediately consuming the food items. The rats unloaded their cheek pouches at burrow entrances and pushed the seeds into the burrow with their forelimbs. The Texas kangaroo rat was not observed scatter hoarding seeds as does D. merriami (Daly et al. 1992). Foraging kangaroo rats sometimes covered considerable distances in a single bound and often jumped over taller vegetation. Foraging rats occasionally jumped up and backward during their foraging sessions. Similar backward leaping has been reported in D. deserti, D. ingens, and D. spectabilis as an avoidance mechanism against snakes (Randall 1993; 2007), however no snakes were observed near foraging D. elator. Movements of foraging cattle near the kangaroo rats caused them to briefly enter their burrows. Howling coyotes (Canis latrans) caused the kangaroo rats to stop foraging, freeze, and sometimes rise up on their hind legs. Although some burrows were located 2 m from an adjacent wheat field, foraging always occurred in the pasture and never in the wheat field. No kangaroo rats were observed within the wheat field area either before or after harvesting of the grain. Foraging distances were obtained for seven individuals. One individual foraged a distance of 2.5 m from its burrow, another 5 m from its burrow, four individuals foraged 15m from their burrows, and one ranged 20 m from its burrow. The longest movement occurred when a kangaroo rat left one burrow and traveled 26 m directly to another burrow without foraging. Roberts and Packard (1973), using incandescent lighting, reported maximum distances of 314 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 86.89 m for males and 108.53 m for females and found that some D. elator traveled more than 300 m along roads at night. Because infrared rather than incandescent light was used during this study, the observations of shorter foraging distances may be more indicative of the Texas kangaroo rat's foraging behaviors. Temporal, thunderstorm and moonlight effects on foraging behaviors -OXhQX researchers noted that the Texas kangaroo rat was most active two to three hours after darkness (Packard & Roberts 1973; Carter et al. 1985). Initial activity of Texas kangaroo rats during this study began at one hour or less after darkness. Active kangaroo rats were also observed during early morning hours (330 and 400 hrs). In one case, Texas kangaroo rats were observed from 2230 hrs to sunrise. No differences in activity levels or foraging behavior were noticed during any of these periods of time. Texas kangaroo rats continuously foraged as lightening from a nearby thunderstorm illuminated the pastureland. No precipitation occurred during this event. During another early morning storm that had high winds and frequent lightening and thunder, a kangaroo rat that had been foraging about 3 m from its burrow entrance ducked back into its burrow after each lightening flash. The kangaroo rat repeated this behavior five times over a period of 20 minutes. Once precipitation began, the rat entered and remained within its burrow until after the storm moved out of the area. Other researchers have stated that the Texas kangaroo rat is not active during moonlit periods (Dalquest & Homer 1984; Jones et al. 1988), or that moonlight foraging is unusual for the species (Packard & Roberts 1973). During this study, Texas kangaroo rats were observed foraging during new, crescent, half, and full moon phases. Randall (1993) observed that heteromyid rodents shift their activities from open areas to more protected bmsh and tree covered areas during moonlit nights. TEXAS J. SCI. 60(4), NOVEMBER, 2008 315 Burrow utilization -Ddlc^QsX & Collier (1964) and Schmidly (2004) stated that Texas kangaroo rat burrows were left unplugged during the day. Of five burrows known to be occupied by D. elator during the night, three were found plugged the following morning. Observational data from this study suggests that D. elator may occupy more than one day burrow. Runways were found connect¬ ing burrows, and observations of animals leaving one burrow and entering and remaining within an adjacent burrow were documented on three separate occasions (21, 22, and 25 May 2005). Trapping data obtained during an earlier study (Goetze et al. 2007) also supports this hypothesis. Packard & Roberts (1973) reported no use of multiple burrows based only on trapping data. Behrends et al. (1986) reported that D. merriami utilized two or more day burrows and occasionally two different individuals may occupy the same burrow at different times. Acknowledgments We wish to thank Mr. and Mrs. Oscar Goetze and Mr. Ernest Goetze for allowing us access to their properties within Wichita County. Our study could not have been completed without their kind permission. We acknowledge Margaret Goetze for field assistance with this project. We thank Brian R. Chapman and other anonymous reviewers for their reviews and corrections which greatly improved earlier versions of this manuscript. This study was conducted under Texas Parks and Wildlife permit no. SPR- 0496-775. Literature Cited Allison, N. 1. & S. Destefano. 2006. Equipment and techniques for nocturnal wildlife studies. Wildlife Society Bulletin, 34(4): 1036-1044. Behrends, P., M. Daly & M. I. Wilson. 1986. Range use patterns and spatial relationships of Merriam's kangaroo rats {Dipodomys merriami). Behaviour, 96(3): 187-209. Carter, D. C., W. D. Webster, J. K. Jones, Jr., C. Jones & R. D. Suttkus. 1985. Dipodomys elator. Mammalian Species, 232:1-3. 316 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Chapman, B. R. 1972. Food habits of Loring's kangaroo rat, Dipodomys elator. Journal of Mammalogy, 53(4):877-880. Dalquest, W. W. & G. Collier. 1964. Notes on Dipodomys elator, a rare kangaroo rat. The Southwestern Naturalist, 9(3): 146-150. Dalquest, W. W. & N. V. Homer. 1984. Mammals of north-central Texas. Midwestern State University Press, Wichita Falls, Texas, 261 pp. Daly, M., L. F. Jacobs, M. 1. Wilson & P. R. Behrends. 1992. Scatter hoarding by kangaroo rats {Dipodomys merriami) and pilferage from their caches. Behavioral Ecology, 3(2): 102-1 11. Goetze, J. R., W. C. Stasey, A. D. Nelson & P. Sudman. 2007. Habitat attributes and population size of Texas kangaroo rats on an intensely grazed pasture in Wichita County, Texas. Texas Journal of Science, 59(1): 1 1-22. Jones, C., M. A. Bogan & L. M. Mount. 1988. Status of the Texas kangaroo rat {Dipodomys elator). Texas Journal of Science, 40(3):249-258. Jones, W. T. 1993. The social systems of heteromyid rodents. Pp. 575-595 in Biology of the heteromyidae (H. H. Genoways & J. H. Brown eds.), 719 pp. Packard, R. L. & J. D. Roberts. 1973. Observations on the behavior of the Texas kangaroo rat. Mammalia, 55(4):680-682. Randall, J. A. 1989. Neighbor recognition in a solitary desert rodent {Dipodomys merriami). Ecology, 8 1(2): 123- 133. Randall, J. A. 1991. Sandbathing to establish familiarity in the Merriam’s kangaroo rat, Dipodomys merriami. Animal Behavior, 41(2):267-275. Randall, J. A. 1993. Behavioural adaptations of desert rodents (Heteromyidae). Animal Behavior, 45(2):263-287. Randall, J. A. 2007. Environmental constraints and the evolution of sociality in semifossorial desert rodents. Pp. 368-379 in Rodent societies: An ecological & evolutionary perspective (J. O. Wolff & P. W. Sherman eds.), 610 pp. Roberts, J. D. & R. L. Packard. 1973. Comments on movements, home range and ecology of the Texas kangaroo rat, Dipodomys elator Merriam. Journal of Mammalogy, 54(4):957-962. Schmidly, D. J. 2004. The mammals of Texas, revised edition. University of Texas Press, Austin, 501 pp. Stangl, F. B., Jr., T. S. Schafer, J. R. Goetze & W. Pinchak. 1992. Opportunistic use of modified and disturbed habitat by the Texas kangaroo rat {Dipodomys elator). Texas Journal of Science, 44(l):25-35. Stangl, F. B., Jr., J. R. Goetze, M. M. Shipley & D. A. Early. 2006. Characterization of the dorsal skin gland of the Texas kangaroo rat, Dipodomys elator (Rodentia: Heteromyidae). Western North American Naturalist, 66(4):462-472. Yoemg, S. I. 1999. Solitary is not asocial: effects of social contact in kangaroo rats (Heteromyidae: Dipodomys heermanni). Ethology, 105(4):3 17-333. JRG at: jgoetze@laredo.edu TEXAS J. SCI. 60(4), NOVEMBER, 2008 317 REPRODUCTIVE CYCLE OF THE BROWN FOREST SKINK, SPHENOMORPHUS CHE RRIEI (SQU AM AT A: SCINCIDAE), FROM COSTA RICA Stephen R. Goldberg Department of Biology, Whittier College, PO Box 634 Whittier, California 90608 The brown forest skink, Sphenomorphus cherriei is an oviparous forest floor inhabitant known from central Veracruz, Mexico to extreme western Panama (Fitch 1983; Savage 2002). In Costa Rica it is an abundant inhabitant of the leaf litter community (Guyer & Donnelly 2004). Previous reports on S. cherriei reproduction are in: (Greene 1969; Fitch 1970, 1973a,b; Alvarez del Toro 1982; Kohler 2000; Savage 2002; Watling et al.2005). The purpose of this paper is to provide additional information on reproduction of S. cherriei from a histological examination of gonadal material from museum specimens as part of an ongoing study on the reproductive biology of Costa Rican lizards. The first information on the testicular cycle and evidence that S. cherriei produces multiple clutches are presented. Specimens of S. cherriei from Costa Rica collected 1959 to 1986 were examined from the herpetology collection of the Natural History Museum of Los Angeles County (LACM), Los Angeles, California. Thirty three males (mean SVL = 49.6 mm ± 4.6 SD, range = 41-57 mm); fifty-three females (mean SVL = 53.4 mm ±4.8 SD, range = 36- 64 mm) and seventeen presumably neonates (mean SVL = 22.2 mm ± 1.4 SD, range = 18-23 mm) were examined. The left testis and epididymis were removed from males and the left ovary was removed from females for histological examination. Enlarged follicles (> 4 mm) or oviductal eggs were counted; no histology was performed on them. Tissues were embedded in paraffin, sectioned at 5 pm and stained with Harris’ hematoxylin followed by eosin counterstain (Presnell & Schreibman 1997). An unpaired /-test was used to compare S. cherriei male and female mean body sizes using non- parametric data and the relationship between female SVL and clutch size was examined by linear regression analysis using Instat (vers. 3.0b, Graphpad Software, San Diego, CA). 318 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Material examined. -ThQ following specimens of S. cherriei from Costa Rica (by province) were examined: Alajuela (1): LACM 161700. Cartago (7): LACM 161672, 161673, 161868, 161913, 161914, 162562, 162598. Guanacaste (23): LACM 161615, 161619, 161667, 161691, 162565, 162567, 162571, 162573, 162575, 162580, 162582, 162590, 162592, 162595, 162596, 162601, 162610, 162614, 162616, 162620, 162621, 162623, 162628. Heredia (25): LACM 161618, 161632, 161659, 161690, 161865, 161869, 161878, 161880, 161682, 161694, 161869, 161870, 161882, 161884, 161886, 161895, 161912, 161916, 161917, 161929, 161934, 162586, 162600, 162603, 162612. Limon (22): LACM 161617, 161633, 161644, 161645, 161661, 161675, 161681, 161693, 161872, 161874, 161894, 161899, 161901, 161902, 161903, 161905, 161906, 161910, 161911, 161918, 162569, 162608. Puntarenas (23): LACM 161628, 161629, 161636, 161650, 161653, 161656, 161657, 161671, 161676, 161684-161686, 161692, 161695, 161705, 161876, 161907, 162557, 162560, 162561, 162564, 162599, 162625. San Jose (2): LACM 161665, 162572. Stages in the testicular cycle are presented by month in Table 1. Three stages were present: (1) Recrudescent: (renewal) in which there is a proliferation of germinal cells, spermatogonia and primary spermatocytes predominate; (2) Late Recrudescent: numbers of germinal cells have increased; primary, secondary spermatocytes, spermatids, but no spermatozoa are present; (3) Spermiogenesis, clusters of sperm line the seminiferous tubules, metamorphosing sperm are present; epididymidis contains sperm. Males undergoing spermiogenesis were present in each month (Table 1) indicating a continuous testicular cycle. The smallest reproductively active male (LACM 161886) measured 41 mm SVL and was collected in April. In another reproductive study on a lizard from Costa Rica, Marion & Sexton (1971) reported a seasonal testicular cycle for Sceloporus malachiticus from high elevations (800 to 3200 m). In November- January (dry season) testes were in regression (spermatogonia predominate) or recrudescent; by June maximum spermiogenesis was in progress (Marion & Sexton 1971). Females of S. cherriei were significantly larger than males (unpaired t test, t = 3.1, M df, P = 0.0004). Reproductively active TEXAS J. SCL 60(4), NOVEMBER, 2008 319 Table L Monthly distribution of reproductive conditions in the testicular cycle of 33 Sphenomorphus cherriei from Costa Rica. Values are the numbers of males exhibiting each of the conditions. Month N Recrudescent Late Recrudescent Spermiogenesis February 1 0 0 1 April 5 0 0 5 May 7 0 1 6 June 3 0 0 3 July 7 1 0 6 August 3 0 1 2 September 2 0 0 2 October 1 0 0 1 November 3 0 0 3 December 1 0 0 1 Table 2. Monthly distribution of reproductive conditions in the ovaries of 53 Sphenomorphus cherriei from Costa Rica. Values shown are the numbers of females exhibiting each condition. Month n No yolk deposition Early yolk deposition Enlarged follicles >4mm Oviductal eggs Oviductal eggs and yolk deposition January 2 0 1 1 0 0 March 6 0 0 3 2 1 April 6 1 0 1 3 1 May 9 1 3 3 2 0 June 7 0 2 1 2 2 July 12 2 2 3 5 0 August 6 2 1 1 2 0 September 3 0 1 1 0 1 October 1 0 0 1 0 0 December 1 0 1 0 0 0 females were present in each month (Table 2) indicating a continuous ovarian cycle. Five females (Table 2) with oviductal eggs were undergoing concomitant early yolk deposition (= basophilic yolk granules in the ovarian follicle) for a subsequent clutch. This is the first definitive evidence that S. cherriei produces multiple clutches. Mean clutch size (n = 36) was 2.3 ± 0.65 SD, range =^1-3. Linear regression analysis revealed a significant positive correlation between female body size (SVL) in mm and clutch size for 36 cherriei 320 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 females: Y = - 0.01 + 0.04X, r = 0.35, P = 0.04. The smallest reproductively active female (collected in May) measured 46 mm (LACM 161673), with 2 follicles > 4 mm. My estimation of minimum size for female reproductive activity of S. cherriei of 46 mm is close to that of Greene (1969) who reported 44 mm. Sphenomorphus cherriei of neonate size (18-23 mm) were collected in March, April, May, June, July and August indicating an extended reproductive season. Fitch (1973a,b) who analyzed samples from Turrialba, Cartago Province, Costa Rica found most females were non-reproductive during the dry season (January to April) indicating there was some seasonality in S. cherriei reproduction. One January female from Cartago Province (LAM 162562) was undergoing early yolk deposition, however, it is not known when it would have been completed. According to Fitch (1973b; 1983) S. cherriei reproduces throughout the year in the humid Caribbean lowlands. Alvarez del Toro (1982) reported S. cherriei in Chiapas produced clutches of 3-4 eggs in June and July (maximum clutch size reported herein is 3) and Fitch (1985) reported the possibility of geographic variation in clutch sizes. Examination of additional gonadal samples from different populations is required before the degree of geographic variation in the reproductive cycle of S. cherriei can be ascertained. Acknowledgments I thank Christine Thacker (Natural History Museum of Los Angeles County) for permission to examine S. cherriei and Sean Kark (Whittier College) for assistance with histology. Specimens are part of the CRE (Costa Rica Expeditions) collection donated to LACM by Jay Savage in 1998. Literature Cited Alvarez del Toro, M. 1982. Los reptiles de Chiapas, 3""^ Edic., Instituto de Historia Natural del Estado, Tuxtla Gutierrez, Chiapas, Mexico, 248 pp. Fitch, H. S. 1970. Reproductive cycles in lizards and snakes. Univ. Kansas Mus. Nat. Hist., Misc. Publ. No. 52:1-247. Fitch, H. S. 1973a. A field study of Costa Rican lizards. Univ. Kansas Sci. Bull. 50:39- 126. TEXAS J. SCI. 60(4), NOVEMBER, 2008 321 Fitch, H. S. 1973b. Population structure and survivorship in some Costa Rican lizards. Occas. Pap. Mus. Nat. Hist., Univ. Kansas, 18:1-41. Fitch, H. S. 1983. Sphenomorphus cherriei (Escincela Parda, Skink). Pp. 422-425 in D.H. Janzen, editor, Costa Rican Natural History. The University of Chicago Press, Chicago, 816 pp. Fitch, H. S. 1985. Variation in clutch and litter size in New World reptiles. Univ. Kansas, Mus. Nat. Hist., Misc. Publ. No. 76:1-76. Greene, H. W. 1969. Reproduction in a Middle American skink, Leiolopisma cherriei (Cope). Herpetologica, 25:55-56. Guyer, C. & M. A. Donnelly. 2004. Amphibians and Reptiles of La Selva, Costa Rica, and the Caribbean Slope: A comprehensive guide. University of California Press, Berkeley, viii + 299 pp. Kohler, G. 2000. Reptilen und amphibien Mittelamerikas. Band 1: krokodile, schildkroten, echsen. Herpeton, Offenbach, Germany, 159 pp. Marion, K. R. & O. J. Sexton. 1971. The reproductive cycle of the lizard Sceloporus malachiticus m Copeia, 1971:517-526. Presnell, J. K. & M. P. Schreibman. 1997. Humason’s Animal Tissue Techniques, 5* Ed. Johns Hopkins Press, Baltimore, xix + 572 pp. Savage, J. M. 2002. The amphibians and reptiles of Costa Rica, A herpetofauna between two continents, between two seas. The University of Chicago Press, Chicago. 934 pp. Watling, J. I., J. H. Waddle, D. Kizirian & M. A. Donnelly. 2005. Reproductive phenology of three lizard species in Costa Rica, with comments on seasonal reproduction of neotropical lizards. J. Herpetol., 39:341-348. SRG at: sgoldberg@whittier.edu THE DISTRIBUTION OF SPOTTED SKUNKS, GENUS SPILOGALE, IN TEXAS Robert C. Dowler, Carla E. Ebeling, Gema I. Guerra and Adam W. Ferguson Department of Biology, Angelo State University San Angelo, Texas 76909 Spotted skunks (genus Spilogale) are small skunks that occur over much of the continental United States and Mexico, ranging north to British Columbia and south to Costa Rica (Hall 1981). The taxonomic history of members of the genus in the contiguous 48 states has ranged from as many as 10 species (Howell 1906) to their synonymy into one polytypic species, S. putorius, by Van Gelder (1959). In Texas, early editions of The Mammals of Texas followed 322 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 this pattern with recognition of three species of spotted skunks (Taylor & Davis 1947) followed by only one (Davis 1966). Research by Mead (1968a; 1968b) revealed that eastern and western forms of S. putorius had distinctly different breeding times, with western forms breeding in the early fall and having delayed implantation, whereas eastern forms breed in the spring with no delayed implantation. Mead (1968b) argued that this reproductive pattern acted as a temporal reproductive isolating mechanism and suggested the eastern and western spotted skunks be treated as two species, S. putorius and S. gracilis^ respectively. Since that time, several mammalian classifications continued to recognize only S. putorius in the United States (Hall 1981; Corbett & Hill 1991; Wozencraft 1993) but the most recent carnivore classification treated both eastern and western spotted skunks as different species (Wozencraft 2005). In Texas, editions of The Mammals of Texas since 1974 (Davis 1974; Schmidly & Davis 1994; Schmidly 2004) have followed such a classification recognizing the existence of both species in the state. Although the taxonomic status of spotted skunks in Texas appears resolved, the current distributional ranges of these two species remain unclear. All depictions of the distribution of the two species of spotted skunks in Texas have left the central part of the state without records and presumably the two did not meet in much of this part of the state (Schmidly 2004). Counties without records (Schmidly 2004) include Nolan, Taylor, Callahan, Eastland, Runnels, Coleman, Brown, Comanche, McCullough, San Saba, Mills, Hamilton, Lampasas, and Burnet. Recent recovery of records, reported herein, for seven counties in west-central Texas clarify the distribution of spotted skunks in Texas (Fig. 1). Specimens are deposited in the Angelo State Natural History Collections (ASNHC), Angelo State University and the Texas Cooperative Wildlife Collections (TCWC), Texas A&M University. TEXAS J. SCI. 60(4), NOVEMBER, 2008 323 ^\;PALO PINTO, ’B0RDEN#'SCURRY s\FISHER<\\ JONES^ GAINES DAWSON EASTLAND- "^ANDREWS TWrtin"'^;' HOWARD'! Mitchell"^ nolan TAYLOR erath: CALLAHAN 'COMANCHE' RUNNELS REAGAN 'MCCULLOUGH ■sansaba>laivipasas. SCHLEICHER . MENARD ?MASON^LLANO- SUTTON^ -'z ^ , .■ ;haYS\ //'EDWARDS T'' ' A z V/s^^OVC^KENC w/^<^Vz;/zV//v^ Y/ KINNEY ;/T,;, UVALDE;' v0MEDINA/ Figure 1. Range map of Spilogale gracilis [^] and S. putorius [^] in Texas adapted from Schmidly (2004) identifying the area in central Texas (white) where Spilogale was thought to be absent (inset). Area of potential range overlap between the two species is identified by ® . Map indicating records of S. gracilis [•] and S. putorius [A] from central Texas with county records reported herein outlined in bold. The Erath County record was previously reported in Goetze et al. (2004). On 21 March 2003, a road-killed specimen (ASNHC 13370) of Spilogale putorius was recovered from Coleman Co., 1.6 km E Valera (UTM 14R 449697, 312919). This animal had an intermediate pattern of pelage spots and stripes between typical S. gracilis and S. putorius; however, white portions of the tail were the same as 5. putorius from east Texas. The specimen was an adult male with testes measuring 14 mm. Body measurements (in mm) were: total length-470, tail length-175, hind foot length-46, and ear length-24; mass was 0.6 kg. Another specimen of S. putorius was donated to Howard Payne University in Brownwood and subsequently transferred to the 324 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Angelo State Natural History Collections (ASNHC 13369). This animal was salvaged 1.6 km NW Brownwood on Route 84 and was taken on 22 February 2004. It was a scrotal male with testes measuring 24 mm. Measurements of this skunk (as above) were 490-168-47-22 and its mass was 650 g. The pattern of stripes and spots was somewhat intermediate between the two species, but was generally typical of eastern spotted skunks. A road-killed specimen (ASNHC 13372) of S. putorius was recovered in Taylor County, 6.9 km N, 3.7 km E Tuscola on 23 April 2007. This animal, also a scrotal male with testes measuring 20 mm, had body measurements of 469-158-45-23 and a mass of 0.8 kg. Its pelage also was intermediate in striping pattern between S. gracilis and S. putorius, but shared the pattern of little white on the ventral side of the tail, typical of S. putorius. A road-killed specimen of S. gracilis (TCWC 59610) was collected on 27 July 2008 on Interstate 10 in Crockett County, 18.2 km west of Ozona (14 R 270901, 3397611) at 711 m elevation. This western spotted skunk was a male measuring 460-150-46-27 with a mass of 726 g. In addition to the specimens described above, two photographs verifying records of western spotted skunks, S. gracilis, in Gaines and Comal counties have come to the attention of the authors. The first photograph is of a Gaines County animal which was killed in a house 1.6 km S Seminole (UTM 13S 0721230, 3619012) on 2 February 1994. This record is at the northern limit of the range of S. gracilis at the southern extreme of the Texas panhandle (Fig. 1). A second photograph is of a western spotted skunk that was trapped and released about 24 km N of New Braunfels in Comal County, (UTM 14R 581695 3310470). This record (Fig. 1) extends the range of the western spotted skunk about 16 km east of its range as depicted by Schmidly (2004). Both photographs have been archived at the ASNHC. TEXAS J. SCI. 60(4), NOVEMBER, 2008 325 Goetze et al. (2004) reported an additional county record for Erath County, Texas, which is housed in the mammal collection at Tarleton State University (TSU 1264). This specimen was collected 9.6 km WNW Stephenville on 10 November 1971. Its gender was unknown. All records, excluding Goetze et al. (2004), are new for counties in Texas, based on Schmidly (2004), and four are range extensions. The records of eastern spotted skunks in Brown, Coleman, and Taylor counties fill much of the gap (Fig. 1) in the reported range of spotted skunks in central Texas (Schmidly 2004). Although these records may indicate recent range expansions, it is just as likely that these represent historical occupation by Spilogale putorius in this region and the species simply has not been detected previously. Compilation of these Spilogale records indicates that the entire state is occupied by at least one species and areas of historical range overlap still remain. Additional monitoring of this genus in Texas should further help increase the understanding of its distribution and conservation status across the state. Acknowledgments Thanks to T. Collins, J. B. Doty, D. J. Leavitt, and J. Spencer for salvaging specimens reported herein. Thanks also to J. L. Strickland and J. R. Ott for photographic records. A special thanks to Dr. Edward Roth, Howard Payne University, for donation of one specimen of Spilogale putorius to the Angelo State Natural History Collections. H. L, Prestridge kindly provided information on the specimen at the TCWC. Lastly, we appreciate all of the students and faculty who have assisted in the salvage and preparation of skunk specimens. Literature Cited Corbet, G. B. & J. E. Hill. 1991. A world list of mammalian species ed.). Nat. Hist. Mus. Pub., Oxford Univ. Press, v + 243 pp. Davis, W. B. 1966. The mammals of Texas. Bull. Texas Game and Fish Commis., 41:1-267. 326 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Davis, W. B. 1974. The mammals of Texas. Bull. Texas Parks and Wildlife Dept., 41:1-294. Goetze. J. R., P. D. Sudman, & A. Nelson. 2004. Noteworthy records of mammals from Erath County, Texas. Occas. Pap. Mus., Texas Tech Univ., 234:1-6. Hall, E. R. 1981. The mammals of North America. 2 vols. John Wiley & Sons, NY, 1181 pp. Howell, A. H. 1906. Revision of the skunks of the genus Spilogale. N. Am. Fauna, 26:1-155. Mead, R. A. 1968a. Reproduction in eastern forms of the spotted skunk (genus Spilogale). J. Zook, 156:119-136. Mead, R. A. 1968b. Reproduction in western forms of the spotted skunk (genus Spilogale). J. Mamm., 49(3):373-390. Schmidly, D. J. 2004. The mammals of Texas (Revised Edition), Univ. of Texas Press, Austin, xx + 501 pp. Schmidly, D. J. & W. B. Davis. 1994. The mammals of Texas. Univ. of Texas Press, Austin, 352 pp. Taylor, W. P. & W. B. Davis. 1947. The mammals of Texas. Bull. Texas Game Fish Oyster Comm., 27:1-79. Van Gelder, R. G. 1959. A taxonomic revision of the spotted skunk (genus Spilogale). Bull. Am. Mus. Nat. Hist., 1 17:229-392. Wozencraft, W. C. 1993. Order Carnivora. Pp. 279-348 in Mammal species of the world: a taxonomic and geographic reference, 2^^ edition. (D. E. Wilson and D. M. Reeder, eds.). Smithsonian Institution Press, Washington, xvii + 1207 pp. Wozencraft, W .C. 2005. Order Carnivora. Pp. 532-628 in Mammal species of the world: a taxonomic and geographic reference. 3*^^ edition. (D. E. Wilson and D. M. Reeder, eds.). The Johns Hopkins University Press, Baltimore, xxxv + 2142 pp. RCD at: robert,dowler@angelo.edu THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 327 INDEX TO VOLUME 60 (2008) THE TEXAS JOURNAL OF SCIENCE Jeffery A* Masters and Cherise D. Montgomery Department of Biology, Angelo State University San Angelo, Texas 76909 ^ This index has separate subject and author sections. Words, phrases, locations, proper names and the scientific names of organisms are followed by the initial page number of the articles in which they appeared. The author index includes the names of all authors followed by the initial page number of their respective article(s). SUBJECT INDEX A Abilene Christian University 189 Acanthophora spicifera 221 Africa, southern 151, 163 Agardhieila ramosissima 22 1 Algal community structure 201 American College Testing (ACT) Program 189 amphibians 123 Analysis of Similarity 83 Anaxyrus speciosus 171 Angelo State University 189 angiosperms 113 Anotrichium sp. 201 anurans 123 Arkansas Gulf Coastal Plain 23 Mississippi Alluvial Plain 23 Arkansas River 23 AutoCAD© 137 B bantam sunfish 23 bill morphology 2 1 5 binary logistic regression 1 89 blue crab 27 1 bluefm killfish 69 Brazos water snake 63 brown alga 227 Bufo nebulifer 123 C Calculus I 1 89 Callinectes sapidus 271 Camp Mabry 123 Capulin Volcano Field 157 Centrarchidae 23 Centroceras sp. 201 Ceramiales 201,221 Ceramium sp. 201 328 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 Chaetoceros sp. 281 Chelydra serpentina 123 chemical oxygen generators 299 Cladophora sp. 3 coastal plain toad 171 Colubridae 63 common barking gecko 1 5 1 common rough-scaled lizard 163 contact angles 137 continental shelf, outer 20 1 Coscinodiscus sp. 281 D Dactyliosolen sp. 281 Dictyotales 227 Digenea 63 Diidomethane 137 Dipodomys elator 309 Ditylum sp. 281 Dixonius siamensis 233 Drop-Ex Plus 299 E electrophoresis, denaturing gradient gel(DGGE) 33 ethylene glyeol 137 explosives, homemade 299 ex-post facto design 1 89 F Flower Garden Banks 20 1 foraging behavior 215 G Gekkonidae 1 5 1 , 233 giant walking stick 57 Gigartinales 221 gland expression 1 7 1 glass microscope slides 137 global climate change 179 gonads 163 gray fox 225 green sunfish 179 Guinar diasp. 281 Gulf of Mexico 83,201,227 northwestern 201 H herpetofaunal inventory 123 heteroduplexes 33 hydrogen peroxide 299 Hypoglossum sp. 201 I Ichnotropis squamulosa 163 J Justicia americana 3 K Kruger National Park 1 5 1 L Lacertidae 163 Lamar University 1 89 Lepomis cyanellus 179 Lepomis symmetricus 23 Lithodesmium sp. 281 Little Karoo 1 5 1 lizards 123 Lucania goodei 69 M macrophytes 3 mandibular structure 215 mark-recapture methodology 45 masked duck 215 mating behavior 57 Megaphasma dentricus 57 Mexieo Coahuila 243 Microlophus koepckeorum 261 Microlophus occipitalis 261 Microlophus peruvianus 26 1 Microlophus stolzmanni 26 1 Microlophus theresiae 26 1 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 329 Microlophus thoracicus 261 Microlophus tigris 261 Mississippi River 23 Montezuma bald cypress 243 Multivariate statistic techniques 83 N Naviculasp. 281 Nerodia harteri 63 New Mexico northeastern 157 Nomonyx dominicus 215 N-type silicon wafers 137 nitrogen-fixing bacteria 97 North Bosque River 3 O Ochetosoma aniarum 63 Ochetosomatidae 63 Ochrophyta 227 Odontellasp, 281 Ollotis nebulifer 171 Olmstead-Tukey diagrams 243 Orthoptera 57 oxidizers 299 Ouachita River 23 Oxyura jamaicensis 215 P Padina glabra 227 painted bunting 243 paratoid glands 171 PCR 33 PCR-RFLP 97 Perciformes 23 periphyton 3 Phaseolus vulgaris 97 Phasmidae 57 photogrammetric samples 201 phytoplankton 3,281 Pinto bean 97 piny on-juniper occurrence 1 5 7 planktonic diatoms 281 Polysiphonia sp. 201 potassium chlorate 299 Pseudonitzchia sp. 281 Ptenopus garrulous 151 R radio transmitters 45 Rana catesbeiana 123 red algal mat 20 1 redox indicators 299 Red River 23 reproduction 151,163,233,261 reptiles 123 Rhizobium etli 97 Rhizobium tropici 97 Rhizosolenia sp. 281 Rio Sabinas 243 riparian bird community 243 ruddy duck 2 1 5 S Saline River 23 secretory vesicle 1 7 1 self-assembled monolayer (SAM) 137 Serpentes 63 Siamese leaf-toed gecko 233 snakes 123 snakes, colubrid 123 South America 261 spermiogenesis 151, 163,233,261, 317 Squamata 151,233,261 St. Francis River 23 surface energetic analyses 137 survivorship models 45 Syngnathidae 83 Syngnathus affinis 83 Syngnathus scovelli 83 Syrrophus marnocki 123 330 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 T Taxodium mucronatum 243 Teleostei 83 temperature tolerance 1 7 9 Texas barrier islands 225 coastal 221, 225 Corpus Christ! Bay 83,221 Edwards Plateau 45 Galveston Bay 281 Lower Rio Grande Valley 97 Offatts Bayou 281 north central 3 Rolling Plains 113 southeast 281 southern 97 Tres Palacios Bay 27 1 Texas Cities Austin 123 Mason 45 Texas Counties Wichita County 113 Texas kangaroo rat 309 Texas National Guard 123 Texas pipefish 83 Texas State University 189 Texas toad 171 Thailand 233 toxin production 171 Trachemys scripta 123 transcutaneous electrical stimulation 1 7 1 trophic status 3 Tropiduridae 261 turtles 123 U University of Texas at San Antonio 1 89 Urocyon cinereoargenteus 225 V van Oss-Chaudbury-Good equation 137 W White River 23 white-winged doves 45 wood stork 243 Z Zenaida asiatica 45 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 331 AUTHORS Anderson, J.D. 27 1 Kuklenz, K.D. 299 Baccus, J.T. 45 Back, J.A. 3 Beitinger, T.L. 179 Brister, J.S. 3 Bursey, C.R. 63 Lehman, R.L. 201,221 Maginnis, T.L. 57 Martynova- VanKley, A. 33 Materon, L.A. 97 McAllister, C.T. 23,63 Canales-del Castillo, R. 243 Cobb,G.P. 137 Coimbatore, G. 137 Cool,C.L. 57 Cortemeglia, C. 179 McDaniel, L.N. 137 McDonald, D.L. 271 McDonough, M.M. 123 McFarland, A.M.S. 3 Messina, P.F. 189 Millican, J.S. 3 Dowler, R.C, 321 Muniz, J.L. 57 Ebeling, C.E. 321 Eitniear, J.C. 215 Eubanks, T. 97 Nalian, A. 33 Nelson, A.D. 113,309 Powell, R.L. 171 Fechhelm, R.G. 69 Ferguson, A.W. 123, 321 Fikes,R.L. 221 Forstner, M.R.J. 123 Frey,J.K. 225 Purviance, J.R. 171 Quigg,A. 281 Robison, H.W. 23 Roehrbom, L. 281 Gallaway, B J. 69 Romero, N. A. 137 Goetze, J.R. 113,309 Ruvalcaba-Ortega, L 243 Goldberg, S.R. 151,163,233,261, Rylander, K. 215 317 Gonzales-Rojas, J.L 243 Gonzales, R.H., Jr. 171 Grosso, J.L. 157 Guerra, G.I. 321 Schwertner, T.W. 45 Small, M.F. 45 Soto, M. A. 171 Stasey, C. 309 Syvyk, A. 33 Hacker, W.D. 157 Hill,E.M. 201 Howells, R.G. 69 Hume, M. 33 Tolan, J. 83 Tumlison, R. 23 Williams, D.L. 299 Jones, G.D. 225 Wynne, M.J. 227 332 THE TEXAS JOURNAL OF SCIENCE-VOL. 60, NO. 4, 2008 REVIEWERS The Editorial staff wishes to acknowledge the following individuals for serving as reviewers for those manuscripts considered for publication in Volume 60. Without your assistance it would not be possible to maintain the quality of research results published in this volume of the Texas Journal of Science. Loren Ammerman Chris Hansen Dennis Parmley Jim Armacost Kimberly A. Hays Nelson Passos Keith R. Arnold Toby Hibbitts C.O. Patterson Thomas Arsuffi, Jr. Sharon Jasper Christin Pruett Jim Baccus Michelle Knox Antonietta Quigg Guy Bernard Jeff Kopachena Magaly Rincon- Joel E. Brandt Thomas LaPoint Zachary Greg Broussard Thomas E. Lee, Jr. Jon Scales Carla B. Carr Roy L. Lehman Joel Schmitter Brian Chapman Barney Lipscomb William Shear Jerry Cook Felicia Manau Jianguo Shao William Cook James Masuoka Michael Shipley Donald Davies Dana Mills Richard Simpson Hudson DeYoe Chris McAllister Fred Stevens James R. Dixon Karen McBee Betty H. Stewart Fred Gehlbach Dale McDonald Ned Strenth Jim R. Goetze Allan D. Nelson Joseph Tomasso Steven R. Goldberg Gerald R. North Roy Vogtsberger Ranette Halverson Darlene Occena- Kenneth Wilkins Gutierrez Michael Wynne THE TEXAS JOURNAL OF SCIENCE— VOL. 60, NO. 4, 2008 333 IN RECOGNITION OF THEIR ADDITIONAL SUPPORT OF THE TEXAS ACADEMY OF SCIENCE DURING 2008 PATRON MEMBERS Stephen R. Goldberg Deborah D. Hettinger Don W. Killebrew David S. Marsh Ned E. 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PS Form 3526, September 2007 (Page 2 of 3) THE TEXAS ACADEMY OF SCIENCE, 2008-2009 OFFICERS President: President Elect: Vice-President: Immediate Past President: Executive Secretary: Corresponding Secretary: Managing Editor: Manuscript Editor: Treasurer: AAAS Council Representative: International Coordinator: Raymond C. Mathews, Jr., Texas Water Dev. Board William J. Quirm, St. Edward’s University Benjamin A. Pierce, Southwestern University Hudson R. DeYoe, University of Texas-Pan American Fred Stevens, Schreiner University Diane B. Hyatt, Texas Water Development Board Ned E. Strenth, Angelo State University Frederick B. Stangl, Jr., Midwestern State University John A. Ward, Brooke Army Medical Center James W. Westgate, Lamar University Armando J. Contreras, Universidad Autonoma de N.L. DIRECTORS 2006 Herbert D. Grover, Wayland Baptist University Gary P. Garrett, Texas Parks and Wildlife Department 2007 Renard L. Thomas, Texas Southern University Bob Murphy, Texas Parks and Wildlife Department 2008 Christopher M. Ritzi, Sul Ross State University Andrew C. Kasner, Audubon Texas SECTIONAL CHAIRPERSONS Anthropology: Raymond Mauldin, University of Texas at San Antonio Biomedical: G. Scott Weston, University of the Incarnate Word Botany: Joan Hudson, Sam Houston State University Cell and Molecular Biology: Magaly Rincon-Zachary, Midwestern State University Chemistry and Biochemistry: Benny E. Amey, Jr., Sam Houston State University Computer Science: James McGuffee, St. Edward’s University Conservation Ecology: Cathy Early, University of Mary Hardin Baylor Environmental Science: Kenneth R. Summy, University of Texas-Pan American Freshwater Sciences: Romi Burks, Southwestern University Geosciences: Joseph I. Satterfield, Angelo State University Marine Sciences: Larry D. McKinney, Harte Research Institute Mathematics: Elsie M. Campbell, Angelo State University Physics: David L. Bixler, Angelo State University Science Education: R. Russell Wilke, Angelo State University Systematics and Evolutionary Biology: Allan W. Hook, St. Edward’s University Terrestrial Ecology and Management: Christopher M. Ritzi, Sul Ross State University COUNSELORS Collegiate Academy: David S. Marsh, Angelo State University Junior Academy: Vince Schielack, Texas A&M University PERIODICALS THE TEXAS JOURNAL OF SCIENCE Texas Academy of Science CMB 6252 Schreiner University Kerrville, Texas 78028-5697 ***********AUTO**MIXED ADC 010 638 SMITHSONIAN INSTITUTION LIBRARIES NHB25MRC154 PO BOX 37012 WASHINGTON DC 20013-7012