Phytologia (Jan 19, 2017) 99(1)
1
Geographic variation in pentane extractable hydrocarbons in natural populations of
Helianthus annuiis (Asteraceae, Sunflowers)
Robert P. Adams and Amy K. TeBeest
Baylor-Gmver Lab, Baylor University, 1 12 Main Ave., Gruver, TX 79040
robert_Adams@baylor.edu
Walter Holmes
Biology Department, Baylor University, Box 97388, Waco, TX 76798
Jim A. Bartel
San Diego Botanic Garden, P. O. Box 230005, Encinitas, CA 92023
Mark Corbet
7376 SW McVey Ave., Redmond OR 97756
Chauncey Parker
11643 Norse Ave., Truckee, CA 96161
and
David Thornburg
2200 W. Winchester Lane, Cottonwood AZ 86326
ABSTRACT
Populations of Helianthus anniius, ranging from eastern Oklahoma to coastal southern California,
were sampled and the yields of total hydrocarbons (HC) from leaves determined. The highest yielding
populations were in the Texas Panhandle (6.0 - 7.99%) and the lowest yields were in Camp Verde, AZ,
NM mountains, Redland, OR. and San Diego, CA. Medium-high yields were found in northern UT and
southern ID. Four populations near Waco, TX had large yield differences ranging from 3.6 to 6.2%.
Some native populations were contaminated by germplasm from cultivated sunflowers and these
populations had very low yields (2.6 - 3.6%). Population variability in HC yields varied geographically
and also between nearby populations, suggesting the micro-habitat environments are important as well as
limited genetic population size. The frequency distribution (329 individuals) ranged from LO to 12.63%
yield and showed a skewed, normal distribution, with a tail towards highest yielding plants. The mean
was 5.33%, with the top 5% being lai'ger than 8.7% yield. A very low correlation (r=0.18) was found
between leaf size biomass and % yield implying an opportunity to select for high yields and high biomass
concurrently. Published on-line www.phytologia.org Phytologia 99(1): 1~10 (Jan 19, 2017). ISSN
030319430.
KEY WORDS: Helianthus annuus, Sunflower, geographic variation in leaf hydrocarbon yields.
Adams and Seiler (1984) surveyed 39 taxa of sunflowers for their cyclohexane (hydrocai*bon) and
methanol (resins) concentrations. The highest cyclohexane (bio-crude) yielding taxa were H. agrestis, an
annual, Bradenton, FL (7.38%) and H. annuus^ Winton, OK (7.09%). Adams et al. (1986) screened 614
taxa from the western US for their hydrocarbon (hexane soluble) and resin (methanol soluble) yields.
They reported 2 plants of H. annuus from Idaho with 8.71% and 9.39% hydrocarbon yields.
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Phytologia (Jan 19, 2017) 99(1)
Seiler, Carr and Bagby (1991) reported on 28 Helianthus taxa for their yields of oil, polyphenols,
protein and rubber. The rubber was found to be of lower molecular weight than Hevea rubber, but still
appeared to be useful as a plasticizing additive and for coatings inside pipes and containers. Yields of
natural rubber has recently been reported for H. annuus (Pearson et al. (2010a) that ranged from 0.9% to
1.7% rubber in cultivated sunflower cultivars (Fig. 4, Pearson et al. 2010b).
There does not appear to be any information on geographic variation in the yields of
hydrocarbons for H. annuus. The purpose of this report is to present new information on geographic
variation of the yields of pentane extractable hydrocarbons in native, annual sunflower. This is
continuation of our research on sunflowers (Adams and TeBeest, 2016; Adams, et al. 2016).
MATERIALS AND METHODS
Population locations - see Appendix I.
The lowest growing, non-yellowed, 8 mature leaves were collected at stage R 5. 1-5.3 (Figure 1)
when the first flower head opened with mature rays. The leaves were air dried in paper bags at 49° C in a
plant dryer for 24 hr or until 7% moisture was attained.
Figure 1. Growth stages of wild (H. annuus) sunflowers, Gruver, TX. Note black ants on the bud and
leaves in lower right photo (from Adams et al. 2016). Sunflower growth stages termination is from
Schnetter and Miller(1981).
Leaves were ground in a coffee mill (1mm). 3 g of air dried material (7% moisture) were placed
in a 125 ml, screw cap jar with 20 ml pentane, the jar sealed, then placed on an orbital shaker for 18 hr.
The pentane soluble extract was decanted through a Whatman paper filter into a pre-weighed aluminum
pan and the pentane evaporated on a hot plate (50°C) in a hood. The pan with hydrocarbon extract was
weighed and tared.
Phytologia (Jan 19, 2017) 99(1)
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RESULTS
The yields of hydrocarbons (HC) by population are given in Table 1. The highest yield (8.60%)
was from Gruver, TX followed by Lake Tanglewood, TX (8.47%) in the Texas Panhandle. The lowest
yield was from Woodward, OK (2.62%) and Eagle Nest, NM (2.62%) followed by cultivated sunflowers
(Oslo, TX)(3.20%). The Woodward population had smooth leaves as found in cultivated sunflowers.
The plants and leaves were very large, although the heads were small. It appears that the Woodward
population was a product of crosses between native and cultivated sunflowers and this resulted in the very
low oil yield.
To visualize the variation in HC yields, the means were contour mapped (Fig. 2). Notice that the
highest yields are in the Texas Panhandle. The lowest yields are in the west (EN, AZ, RO) and just off
the caprock, east of the Texas Panhandle (PT, QN). The low yield at the WO (Woodward, OK) is in a
population that is likely of hybrid origin between native and cultivated sunflowers. The southern Idaho -
northern Utah area had medium-high yields. Of interest are the four populations near Waco, TX (MC,
EC, LC, HC) that have 6.2, 5.3, 3.6, and 4.9% yields in a very small area. At this time, it is not known if
Figure 2. Geographic variation in % yields of HC by population. The asterisk (*) at the WO population
indicates that the population is likely of hybrid origin between native and cultivated sunflowers. Note the
low yield from a commercial sunflower field near Oslo, TX (lower left). See text for discussion.
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Phytologia (Jan 19, 2017) 99(1)
the variation in yields is due to genetics or the environment. It is interesting that the correlation between
% yield and leaf weight as only r= 0.18 (highly significantly different from zero, df = 327). But the
correlation accounts for only 3.24% (r^) of the variance. Thus, breeding for both increased % yields of
HC and biomass seems feasible.
The variability of yields by population is mapped in Figure 3. Population variability in HC yields
varied geographically and also between adjacent populations, suggesting the micro-habitat environments
are important as well as limited genetic population size. One of the least variable populations was
Pocatello, ID (POI, Fig. 3). This was a population of perhaps 50 plants, growing next to the sidewalk at
an on-ramp to 115. It seems likely that POI is very inbred. The Brigham City, UT (BU) population, in a
disturbed vacant lot where a new mall was recently built, was much more variable (Fig. 3). BU contained
perhaps 100 plants, but a more extensive group of sunflowers grew nearby.
Clearly the most unusual situation was the Waco, TX area where 4 nearby populations (MC, FC,
LC, HC, Fig. 3) showed very small to large amounts of variation in their HC yields. MT (Montrose, KS)
was from only 3 cultivated plants raised from seed, so its small variability may be just chance.
Figure 3. Population variability (coefficient of variation in HC yields) for the 29 populations sampled.
The size (diameter) of the circles is proportional to their coefficient of variation.
Phytologia (Jan 19, 2017) 99(1)
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The total yields of HC per the weight of 8 mature leaves is a measure of the grams of HC per
plant (likely larger than if the entire plant were extraeted). In Table 1, the yields range from 0.114 g/ 8
mature leaves (Eagle Nest, NM) to 1.428 g (Gruver, TX). Variation in yields shows (Fig. 4) the highest
yields were in the Texas Panhandle (1.20 g - 1.23 g) and Ellsworth, KS (1.0 g) and Enid OK (0.88 g).
The lowest yields were in the southwestern United States. Note the difference between the San Diego,
large leaves (SL, .43 g) and small leaves (SS, .27 g). These are plants collected from the same
population. Recall that the % yields were quite similar (Table 1, SS, 4.59%; SL, 4.68%).
The four populations near Waco, TX are quite variable and yields ranged from 0.26 g to 0.74 g.
Whether this is due to micro-habitat environments or genetically isolated populations is not known at this
time.
Figure 4. Geographic variation in the HC yields (g/ weight of 8 mature, dried leaves, basis).
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Phytologia (Jan 19, 2017) 99(1)
The frequency distribution (329 individuals, Fig. 5) shows yields ranged from 1.0 to 12.63% with
a skewed, normal distribution, and tailing towards the highest yielding plants (Fig. 5). The mean was
5.33%, with the top 5% being larger than a 8.7% yield. Seed from high yielding plants have been
collected in preparation to examine genetic and environmental factors.
Figure 5. Frequency distribution of HC yields for 329 H. annuus plants. See text for discussion.
This study revealed the range of variation in native sunflowers is quite large, from 1.0 to 12.63%.
It is remarkable to find such a wide range, but indicates the potential of H. annuus to produce copious
amounts of hydrocarbons for use as fuel and in the petro-chemical industry. Many of the highest yielding
plants were severely eaten by grasshoppers and covered with black (sugar) ants, feeding on resin extruded
from the stem, petioles and leaf bracts. It could be that the high yields were responses to insect damage
that induced defense chemicals. The induction of chemical defenses will be examined in a subsequent
study, along with study of the effects of genetics vs. the environment on the production of hydrocarbons.
LITERATURE CITED
Adams, R. P., M. F. Balandrin, K. J. Brown, G. A. Stone and S. M. Gruel. 1986. Extraction of liquid
fuels and chemical from terrestrial higher plants. Part I. Yields from a survey of 614 western United
States plant taxa. Biomass 9: 255-292.
Adams, R. P. and G. J. Seiler. 1984. Whole plant utilization of sunflowers. Biomass 4:69-80.
Adams, R. P. and A. K. TeBeest. 2016. The effects of gibberellic acid (GA3), Ethrel, seed soaking and
pre-treatment storage temperatures on seed germination of Helianthus annuus and H. petiolaris.
Phytologia 98: 213-218.
Adams, R. P., A. K. TeBeest, B. Vaverka and C. Bensch. 2016. Ontogenetic variation in pentane
extractable hydrocarbons from Helianthus annuus. Phytologia 98: 290-297
Pearson, C. H., K. Cornish, C. M. McMahan, D. J. Rath and M. Whalen. 2010a. Natural rubber
quantification in sunflower using automated solvent extractor. Indust. Crops and Prods. 31: 469-475.
Pearson, C. H., K. Cornish, C. M. McMahan, D. J. Rath, J. L. Brichta and J. E. van Fleet. 2010b.
Agronomic and natural rubber characteristics of sunflower as a rubber-producing plant. Indust. Crops
and Prods. 31: 481-491.
Schnetter, A. A. and J. F. Miller. 1981. Description of sunflower growth stages. Crop Sci. 21: 901-903.
Seiler, G. J., M. E. Carr and M. O. Bagby. 1991. Renewables resources from wild sunflowers
(Helianthus spp., Asteraceae). Econ. Bot. 45: 4-15.
Phytologia (Jan 19, 2017) 99(1)
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Table 1. Yields of hydrocarbons (HC) H. annuus, from natural populations. Coefficient of variation
computed as standai'd deviation / mean.
popn id,
sample ids
population sampled
weight
81vs
% yield
corr'd*
Coef. of
variation
Range of
yields
yield
g/8 Ivs
PT PI - PO
14935 Post, TX
8.83
4.36
0.317
0.385
QNQl-QO
14936 Quanali, TX
14.41
4.32
0.238
(2.88,5.62)
0.623
MK Ml -MO
14939 Meade, KS
10.38
5.64
0.186
(3.91,7.21)
0.585
DKDl-DO
14940 Dodge City, KS
9.75
3.52
0.262
(2.68,5.49)
0.346
EKEl-EO
14941 Ellsworth, KS
18.74
5.38
0.240
(3.63,7.35)
1.008
TOTl-TO
14942 Tulsa. OK
13.64
4.56
0.236
3.16,6.04)
0.622
EOOl-OT
14943 Enid, OK
17.60
4.97
0.239
0.875
WO Wl-WO
14944 Woodward, OK,
very large, smooth leaves
19.64
2.62
0.155
(1.92-3.09)
0.515
STSl-SO
14945 grown from seed, ex
Sonora" TX,P1413168
11.47
5.19
0.329
(1.99-7.55)
0.595
OS Ol-TO
6.75
5.95
0.203
(4.19-8.17)
0.406
14947 Lake Tanglewood, TX
■IH
LT: Ll-LO
7/12/16, 1st collection
16.65
(6.73-12.15)
1.410
L2: LA-LJ
7-20-16, 2nd collection
13.92
(5.63-9.06)
1.018
0. 1 95 avg
(5.63-12.15)
1.214
ID 11-19
14948 grown from seed, ex
Idaho, PI 531028
2.77
3.23
0.432
(1.0-6.14)
0.089
SS SA-SJ
14950 San Diego, CA,
small leaves
5.83
4.59
0.292
(2.75-7.51)
0.268
SL SK-ST
14951 San Diego, CA,
large leaves
9.04
4.68
0.218
(2.75-6.11)
0.432
14952 Gruver, TX
GT1:G1-G0
GTl 1-10 1 mi south
16.93
7.26
0.244
(5.01-11.06)
GT2:GA-GJ
GT2 11-20 Imi south
18.03
7.92
0.198
(6.25-10.78)
GT3:GKGT
GT3 21-30,2mi E, Rodeo
12.50
8.16
0.164
(7.00-10.51)
1.020
GT4:A1-AT
GT4 31-40, 1 mi south
14.50
8.60
0.235
(6.52-12.63)
1.247
7.99 avg
0.201 avg
(5.01-12.63)
1.231
SC 10-60
14953 cultivated sunflower
crop, Slough Farm, Oslo, TX
12.41
3.20
0.134
(2.75-3.85)
0.397
MC IM-OM
14976 McLennan Co., TX
Holmes 16654
11.95
6.18
0.144
(4.74-7.76)
0.739
EC IF-OF
14977 Falls Co., Satin, TX
Holmes 16656
9.49
5.29
0.142
(4.19-6.59)
0.502
EC IL-OL
14978 Limestone Co. Mt.
Calm, TX Holmes 16658
6.13
3.58
0.313
(2.61-6.25)
0.219
HC IH-OH
14979 Hill Co., TX Holmes
16661
5.21
4.92
0.372
(2.61-8.65)
0.263
EN lE-OE
14980 Eagle Nest, NM
4.34
2.62
0.326
0.114
LUUl-UO
15023 Logan, UT
7.57
5.44
0.257
(3.98-8.67)
0.412
PI IP-OP
15024 Preston, IT
4.29
6.30
0.278
(3.91-9.34)
0.270
POI II-OI
15025 Pocatello, ID
7.99
5.71
0.160
(4.46-7.55)
0.456
SEC 1 U-OU
15026 Mill Creek, Salt
Lake City, UT
9.54
5.74
0.266
(3.91-8.22)
0.548
RORl-RO
15027 Redmond, OR
5.74
4.06
0.226
(3.37-6.28)
0.233
CN IC-OC
14981 Capuhn, NM
3.51
5.29
0.280
(3.16-8.34)
0.186
MT MA-MC
14982 grown from seed ex
Montrose, KS, PI 413033
8.44
4.91
0.089
(4.65-5.42)
0.414
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Phytologia (Jan 19, 2017) 99(1)
popn id,
sample ids
population sampled
weight
8 Ivs
% yield
corr'd*
Coef. of
variation
Range of
yields
yield
g/8 Ivs
AZ Zl-ZO
15021 Camp Verde, AZ
4.48
3.79
0.332
(1.72-5.56)
0.170
BUBl-BO
15022 Brigham City, UT
5.70
5.90
0.312
(2.90^8.31)
0.336
RNRl-RO
15029 Reno, NV
2.87
5.11
0.299
(2.89-7.90)
0.142
*coiTection factor = soxhlet, 6hr extraction/ pentane 18 hr shaker yield = 2.06
Appendix I Population locations.
Helianlhus petiolaris
common along roadside in sandy soil, flowering. 8.3 mi SW of Fritch, TX on TX 136, 35° 31' 53" N, 101° 38' 31" W. 3360 ft,
Date: 4 June 2016, County: Potter; State: TX
Coll. Robert P. Adams No. 14937
Helianthiis annuiis L. below:
common along railroad and roadside in sandy soil, flowering. 5.3 mi SE of Post TX on US 84, 33° OF 53" N, 101° IT 25" W,
2300 ft. Date: 4 June 2016 County; Garza; State: TX
Coll. Robert P. Adams No. 14935
common along fence row and roadside in sandy soil, flowering.? mi SE of Quanah, TX on US 287,34° 15' 57" N, 99° 36' 46" W,
1450 ft. Date: 5 June 2016 County; Hardeman; State: TX
Coll. Robert P. Adams No. 14936
1.5 mi s of Meade, on KS23, low area in edge of wheat field, 100s of plants in population, but generally uncommon. ~5%
flowering. 37° 15' 49" N, 100° 20' 40" W, 2433 ft. Date; 7 July 2016; County: Meade; State: KS
Coll. Robert P. Adams No. 14939
8.5 mi NE of Dodge City, US 50, several on dirt piles of highway dept., but generally uncommon. ~5% flowering, 37° 47' 06" N,
99° 53' 14" W. 2534 ft. Date; 7 July 2016, County: Ford; State: KS
Coll. Robert P. Adams No. 14940
1.6 mi e of Ellsworth on KS140, on fence row on s side of wheat field, 20 plants, but generally uncommon. -10% flowering. 38°
44' 24" N, 98° 1 1' 53" W, 1600 ft. Date: 7 July 2016, County: Ellsworth; State; KS
Coll. Robert P. Adams No. 14941
15 plants on disturbed area next to South Ash St. (just south of OK364), but generally uncommon, Jenks, OK (sw suburb of
Tulsa). -5% flowering. 36° 00' 57.85" N, 95° 58' 07.61" W, 613 ft. Date: 9 July 2016, County: Tulsa; State: OK
Coll. Robert P. Adams No. 14942
5.5 mi e of Enid on OK412,on fence row, side of wheat field, few plants but generally uncommon, ca 5% flowering, -5%
flowering. 36° 23' 51" N, 97° 46' 51" W, 1160 ft.. Date: 9 July 2016, County: Garfield; State; OK
Coll. Robert P. Adams No. 14943
smooth leaves! 2.8 mi e of Woodward on OK412,on fence row, side of grass field, few plants but generally uncommon, ca 5%
flowering mostly pre-flowering. 36° 25' 53" N, 99° 20' 28" W, 1880 ft.. Date: 9 July 2016, County: Woodward; State: OK
Coll. Robert P. Adams No. 14944
cultivated at Oslo, TX, from seed (USDA P1413168-NC7) ex Sonora, TX. 80% flowering, 36° 25' 12.3" N, 101° 31' 54.6" W,
3239 ft, Date: 12 July 2016, County: cult in Hansford; State: TX.
Coll. Robert P. Adams No. 14945
Phytologia (Jan 19, 2017) 99(1)
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native in grassland, JP & Amy TeBeest farm, 1 mi. s of Oslo Lutheran Church. ~5% flowering. 36° 25' 12.3" N, 101° 31' 54.6"
W, 3239 ft.. Date: 12 July 2016, County: Hansford; State: TX
Coll. Robert P. Adams No. 14946
2- 3 ft plants, lots of resin on petioles and leaf veins, many sugar (black) ants, most with wilted leaves, very dry in July, common
in native grass and on disturbed roadside, brush dump area. Lake Tanglewood, -50% flowering, 35° 04' 23.7" N, 101° 47' 29.0"
W, 3239 ft.. Date: 12 July 2016, County: Randall; State: TX
Coll. Robert P. Adams No. 14947
cultivated at Oslo, TX, from seed (USDA PI 531028) ex Idaho, 80% flowering.36° 25' 12.3" N, 101° 31' 54.6" W, 3239 ft.
Date: 12 July 2016, County: cult in Hansford; State: TX
Coll. Robert P. Adams No. 14948
plants 2' tall, with small leaves, along San Pasqual Rd, 33° 05' 08.2" N, 117° 01' 46.2" W, 353 ft.
Date: 6 July 2016, County: San Diego; State: CA, Coll. Jim A. Bartel 1636
Lab Acc. Robert P. Adams No. 14950
plants to 8' tall, with large leaves, along San Pasqual Rd, 33° 05' 08.2" N, 117° 01' 46.2" W, 353 ft/ Date: 8 July 2016, County:
San Diego; State: CA, Coll. Jim. Bartel 1636
Lab Acc. Robert P. Adams No. 14951
2-3' tall, 10% flowering, lots of damage to leaves by grasshoppers, etc., some with many black (sugar) ants, copious resin at base
of leaves, along fence row, on TX 206, 1-5: 1.2 mi s, 6-10: 1.3 mi. s of Gruver, TX. 36° 14' 52" N, 101° 24' 52" W, 3161 ft.
Date: 16 July 2016, County: Hansford; State: TX
Coll. Robert P. Adams No. 14952
cultivated, irrigated near Oslo, TX, on Slough farm, at R-5.1 stage. 36° 22' 42.17" N, 101° 37' 21.4" W, 3350 ft., leaves mostly
smooth. Date: 17 July 2016, County: cult in Hansford; State: TX
Coll. Robert P. Adams No. 14953
Coll. Walter Holmes
(WCH 16654) McLennan Co. 12* Street at Flat Creek, Robinson (Waco), 27 July 2016 , Walter Holmes
Lab Acc. Robert P. Adams 14976
(WCH 16656) Falls Co. near Satin on FR 434, prairie roadside, 28 July 2016, Walter Holmes
Lab Acc. Robert P. Adams 14977
(WCH 16658) Limestone Co. near jet of Limestone Co roads 102 and 106, south of Mt. Calm, prairie, 29 July 2016, Walter
Holmes
Lab Acc. Robert P. Adams 14978
(WCH 16661) Hill Co. US Hwy 84, West of Mt. Calm near jet with West Somers Lane, 29 July 2016, Walter Holmes
Lab Acc. Robert P. Adams 14979
roadside waste area. Eagle Nest, NM, 36° 33.650' N, 105° 15.969' W, 8260 ft. Date: 8 Aug 2016, County: Colfax; State: New
Mexico, Coll. Amy TeBeest
Lab acc. Robert P. Adams 14980
roadside waste area, Capulin (city), NM, some grasshopper damage, 36° 44.527' N, 104° 00.178' W, 6820 ft.
Date: 8 Aug 2016, County: Union: State: New Mexico, Coll. Amy TeBeest
Lab acc. Robert P. Adams 14981
cultivated at Oslo, TX, from seed (USDA PH413033), ex Montrose, KS. Date: 2 Aug 2016, Coll. Amy TeBeest,
Lab acc. Robert P. Adams 14982
along roadsides. 16-18 mi east of Camp Verde on AZ 260. 34.489° N, 111.597° W, 5900 ft. Date: Aug. 27, 2016, County:
Yavapai; State: AZ, Coll. David Thornburg ns.
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Phytologia (Jan 19, 2017) 99(1)
Lab. acc. Robert P. Adams No. 15021
vacant lot behind new WalMart on disturbed soil, flowering and seeding, multiple branches. W1500S, 775W, Brigham City, UT,
41° 28' 57" N. 1 12° 01' 40" W, 4250 it, Date: Sept. 2, 2016, County; Boxelder; State: UT
Coll. Robert P. Adams No. 15022
vacant lot behind new stores on disturbed soil on US 91 and E2000N, flowering and seeding, multiple branches. Logan, UT, 41°
46' 09" N, 1 1 1° 49' 59" W, 4506 ft. Date: Sept. 2, 2016, County: Cache; State: Utah
Coll. Robert P. Adams No. 1 5023
vacant lot in new subdivision on disturbed soil off of OR hwy 34/36 & just on n edge of Preston, flowering and seeding, multiple
branches. 42° 06' 40" N, 1 1 1 ° 52' 01 " W, 4703 ft. Date: Sept. 2, 2016, County: Franklin; State: Idaho
Coll. Robert P. Adams No. 1 5024
next to sidewalk, on slope, next to freeway (115) access south, flowering and seeding, multiple branches, Pocatello, ID. 42° 52'
49" N, 112°' 25' 35" W, 4625 ft. Date: Sept. 2, 2016, County; Bannock; State; Idaho
Coll. Robert P. Adams No. 1 5025
next to sidewalk, flowering and seeding, multiple branches, common along sidewalks. Mill Creek, UT. s side of 180 on 2000 E,
east side of 2000E. 42° 52’ 49" N, 1 12°' 25' 35" W, 4625 ft. Date: Sept. 3, 2016, County: Salt Lake; State: Utah
Coll. Robert P. Adams No. 1 5026
disturbed ai'ea, vacant on SW Airport Way, ~373m sse of jet SW Airport Way & Veterans Way. Redmond, OR, 44° 15' 30" N,
121°’ 09’ 54" W, 3035 ft, Date; Sept. 3, 2016, County: Redmond; State: Oregon
Coll. Mark R. Corbet, ns, Lab Acc. Robert P. Adams No. 15027
disturbed ai'ea, Neil Rd and west frontage road on 1580, s of Reno, NV. 39° 28' 11.6" N, 119°' 47' 20.4" W, 4485 ft. Date: Sept. 5,
2016, County: Washoe; State: Nevada
Coll. Chaunccy Parker, ns. Lab Acc. Robert P. Adams No. 15029.
Phytologia (Jan 19, 2017) 99(1)
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Seedling growth and leaf photosynthesis of Acer grandidentatum (Bigtooth maple, Sapindaceae)
from isolated central Texas populations
Terri L. Nelson Dickinson
Department of Biology, University of Texas at San Antonio, One UTS A Cirele,
San Antonio, TX 78249, USA terri.nelson.diekinson@gmail.eom
and
O. W. Van Auken
Department of Biology, University of Texas at San Antonio, One UTS A Cirele,
San Antonio, TX 78249, USA osear.vanauken@utsa.edu
ABSTRACT
Two experiments were eompleted to address issues eoneeming apparent reeruitment failure of
native reliet populations of Acer grandidentatum Nutt. (Bigtooth maple) found in the Edwards Plateau
Region of Central Texas. In the first experiment seedlings were grown in pots at 20, 40, 60 and 100%
(open or full sun, 1615 ± 8 pmol/mVs). In the seeond experiment leaf photo synthetie rates of seedlings
growing in sun or shade below a eanopy were examined. Growth of seedlings was greatest at 40% of the
maximum light treatment or at a light level of 705 ± 22 pmol/m /s. Mortality was zero in the 40% light
treatment and 100% at the highest light level tested. Light response eurves were generated using
photosynthetie rates of five leaves of separate juvenile maples growing in full sun or understory eanopy
shade. Rates were measured in the field at light levels from 0-2000 pmol/m /s. From these measurements
a number of photosynthetie parameters were ealeulated and eompared. No signifieant differenees were
seen between the eurves for sun and shade leaves of A. grandidentatum. The only signifieantly different
photosynthetie parameter measured was the maximum photosynthetie rate (Knca)- The A,nax was low at
3.89 ± 0.36 pmol COjIvc^ls for shade leaves and 5.23 ± 0.36 pmol C02/m^/s for sun leaves. The light
saturation point, the light eompensation point and other ealeulated faetors were low as well, but not
signifieantly different. Acer grandidentatum is a shade tolerant speeies with a low photosynthetie rate
whieh seems to be part of the reason it ean persist in isolated Central Texas eanyon woodland
populations. Published on-line www.phytologia.org Phytologia 99(1): 11-21 (Jan 19, 2017). ISSN
030319430.
KEY WORDS: light levels, CO 2 uptake, gas exehange rates, shade plants
It has been ehallenging to understand faetors eontrolling growth and reeruitment of woody
speeies in woodland and forest eommunities, although many researeh papers have dealt with the topie
(Baker et al. 2005). This is true for native reliet populations of Acer grandidentatum Nutt, and other
woody species that are present in the Edwards Plateau Physiographic Region of Central Texas (Van
Auken 1988; Russell and Fowler 2004; Nelson Dickerson and Van Auken 2016). Acer grandidentatum
is a small, deciduous, hardwood tree commonly known as bigtooth maple, but it has many other
common names such as canyon maple, western sugar maple and others (Correll and Johnston 1970;
Tollefson 2006). There has been some debate over the systematics of A. grandidentatum, but most
current papers refer to it simply as A. grandidentatum, which is the convention that we will follow
(Cronquist et al. 1997; Stevens 2001; Atha et al. 2011). The 129 Acer species have been traditionally
grouped into the family Aceraceae, but more recently they have been considered as members of the
subfamily Hippocastanoideae within the family Sapindaceae (Buerki et al. 2009; Watson and
Dallwitz 2011).
Anecdotal reports suggest a decline of recruitment of juvenile A. grandidentatum in central
Texas (Riskind 1979; McCorkle 2007; Adams 2010; BCNPSOT 2010; Heidemann 2011). These
12
Phytologia (Jan 19, 2017) 99(1)
same reports suggest the decline in recruitment is caused by browsing by large herbivores
specifically by Odocoileiis virginianus (white-tailed deer). A recent study confirmed that A.
grandidentatum seedlings protected from large herbivores had a greater rate of survival than
unprotected seedlings (Nelson Dickinson and Van Auken 2016). With leaf removal, a reduction in
plant net photosynthesis occurs. This leaf loss compromises the ability of the plant to replace lost
biomass (Ellswoith et al. 1994)^ which is most readily shown in juveniles. A plant's photosynthetic
parameters affect its inherent growth rate and thus its biomass (Jones and McLeod 1989). Thus,
understanding a plant’s photosynthetic characteristics can help explain how an iudividual plant is
able to compensate for episodes of herbivory and how it adapts to an environment that’s been altered
by herbivoiy (Crowley 1997). Nevertheless, until now, there have been no studies that we could find
concerning photosy nthetic rates of A. grandidentatum,
PURPOSES
The purposes of the present study were to determine the light requirements of juvenile A.
grandidentatum plants and to compare differences in gas exchange rates at various light levels for
leaves of sun and canopy shade A. grandidentatum plants.
METHODS
STUDY AITEAS-There were two study areas. The plant growth experiment was carried out
on a non-shaded roof patio of the science building of the University of Texas San Antonio
(98”34’26”W and 29"37’19”N). The gas exchange measurements were made at Lost Maples State
Natural Area, in Bandera and Real counties about 114 mi west of San Antonio, Texas (29M9’11”N
and 99^^34’59”W).
SEEDLING GROWTH-On March 29, 2010, a total of twenty first-year seedlings were obtained
from a commercial source (Janzow, Boerne, TX), transplanted and randomly placed into one of four light
treatments (five plants/treatment). Plants were randomly placed (one each) into 15.0 cm diameter x 14.5
cm tall pots lined with 3.79 L Ziploc® plastic bags containing 1350 g of soil. Additional nutrients were
added as 5.5 g Osmocote per pot (14/14/14 NPK equivalent to 436 kg/ha nitrogen, 436 kg/ha
phosphorous, and 436 kg/ha potassium). Plants were watered with deionized water as needed, usually 150
mL every day (Janzow 2007). They were placed on a non-shaded roof-top patio on UTSA's Science
building within each of the light treatments.
Plants’ sunlight exposure was limited using shade boxes measuring 0.5 m wide, 0.5 m long, and
1.0 m high. They were constructed with 1.3 cm diameter PVC pipe covered with zero to tliree layers of
commercial black polyethylene shade cloth on five sides secured widi plastic zip ties (Rainbow Gardens
and Lowe's Home Improvement Stores, San Antonio, TX) to adjust light levels in each treatment.
Light levels were measured in each plant location, in each shade box on a clear day in May and
October 2010 within ± 30 minutes of solar noon using a Li-COR® Ll-188 integrating quantum sensor
(Li-COR, Inc. Lincoln, NE). Each shade box contained five A. grandidentatum seedlings (one/pot). Light
levels were 100 % or maximum ( 1615 ± 8 pmol/m^/s, no shade clotli), 60 % (977 ± 42 pmol/mVs), 40 %
(705 ± 22 pmol/nr/s), and 20 % (281 ± 1 pmol/m /s). Boxes were affixed to railings with zip ties and
weighted with sand bags to prevent movement. Pots were covered with clear plastic during rainy weather
to prevent flooding, nutrient and soil loss.
Survival, aboveground, belowground and total dry mass were determined and recorded. Other
plant responses were measured but not reported here (Nelson Dickinson 201 1). Plants were harvested on
October 14, 2010, dried to a constant mass at 80‘^C, and weighed. Data were tested for normality usmg the
Shapiro-Wilks test and for homogeneity of variance using Bartlett's test (Sail et al. 2005). If probability
Phytologia (Jan 19, 2017) 99(1)
13
values fell below 0.05 on either test, data were transformed and retested. Aboveground dry mass,
belowground dry mass, and total diy mass were log transformed, and then analyzed using a one-way
ANOVA followed by Tukey-Kramer HSD. ANOM for proportions was used with a probability level of
0.05 to determine if there were differences in mortality across treatments (McKinley and Van Auken
2005).
GAS EXCHANGE MEASUREMENTS-Ten Acer grandidentatum saplings were randomly
selected in and adjacent to a deer exclosure at Lost Maples State Natural Area, in Sabinal Canyon, Texas.
One fiilly expanded, complete leaf was selected on each plant; five leaves from the shaded canopy
understory plants and five from the open no canopy, full sun plants. The Li-Cor 6400 portable
photosynthetic meter was used to measure gas exchange as a function of light level, or photosynthetically
active radiation (PAR), for each leaf Measurements were made with plants flilly leafed out in May 201 1,
within ± three hours of solar noon using a gas flow rate of 400 pmol/s and a CO 2 concentration of 390
pmol/mol at PARs of 2000, 1600, 1200, 1000, 800, 600, 400, 200, 100, 50, 25, 10, 5, and 0 pmol/niVs.
Each leaf used covered the entire chamber.
Two light response curves were generated, one for sun leaves and one for shade leaves.
Photosynthetic rates along each curve were tested for nonnality using the Shapiro-Wilks test and for
homogeneity of variance using Bartlett’s test (Sail et al. 2005). A repeated measures ANOVA was
completed to deteraiine if there were significant differences between the tu^o leaf types. A one-way
ANOVA with Tukey’s HSD was used to determine differences in photosynthetic rates at different light
levels. For the sun and shade treatments, the maximum rate of photosynthesis (A,,,^.;) was determined,
along with transpiration and conductance at the A,^^- The initial slope of the curve, or quantum yield
efficiency, was also measured. The PAR value at wEich tliis line reached A,„„v was the light saturation
point (L.S/,). Other factors measured were the dark respiration (R^/), the curve's y-intercept and the light
compensation point (L,.,,), the line's x-intercept. These values were also tested for normality using the
Shapiro-Wilks test and for homogeneity of variance using Bartlett's test, then compared using a one-w'ay
ANOVA (Sail et al. 2005).
RESULTS
SEEDLING GROWTH-Total dry mass was significantly different across the four light
treatments (One-way ANOVA; F = 4.6639, P = 0.0159) (Figure 1). The mean total dry mass in the 100 %
sunlight treatment was 0.52 ± 0. 1 1 g/plant. This was significantly different from tlie 40 % treatment, but
was not significantly different from 20 % or 60 % treatments (P = 0.01290, 0.8802, 0.5003 respectively)
(Tukey- Kramer HSD). The mean total dry mass in the 40 % sunlight treatment was greatest at 1.40 ±
0.67 g/plant. This was significantly different from the 100 % treatment {P = 0.0129), marginally different
from the 20 % treatment (P = 0.0555), but not significantly different from the 60 % treatment (P =
0.1335).
Aboveground dry mass was signiifeantly different across the four treatment levels (one-w^ay
ANOVA, P = 0.0156), but only slightly different and only significantly different between the 20 and 100
% treatments (Figure 1). Belowground dry mass was significantly different across all four light treatment
levels (one-way ANOVA, P = 0.0492), but we could not determine where the differences were with the
Tukey-Kramer HSD (multiple range test; Figure 1 ), thus the letters in the figure are all the same across all
light levels.
Mortality was complete in the full sunlight treatment (100 %), while at the 40 % light level, there
was zero mortality (Figure 2). Both of these values were significantly different from the mean at the 0.05
level (ANOM for Proportions; LDL = 0, UDL = 0.868). Twenty percent mortality occurred in the 60 %
14
Phytologia (Jan 19, 2017) 99(1)
light level and 40 % mortality in the 20 % light level, neither of which were statistically significant
(ANOM for Proportions; LDL = 0, UDL = 0.868).
GAS EXCHANGE RATES -Photosynthetic light response curves for full sun and shade leaves of
A. grandidentatum were not significantly different from each other (repeated measures AN OVA, P =
0.0709, Figure 3 A and B). Mean photosynthetic rate for shade leaves of A. grandidentatum was 2.27 ±
0.23 pmol C02/m"/s, which was not significantly different from the mean photosynthetic rate for sun
leaves that was 2.94 ± 0.36 ^unol C02/mVs (one-way ANOVA, P = 0.0709).
Mean maximum photosynthetic rate (A^tv) for shade leaves of A. grandidentatum was 3.89 ± 0.36
pmol C02/m"/s at a PAR of 880 pmol/mVs, while the A,„^a for sun leaves of was 5,23 ± 0.36 pmol
C02/m“/s at a PAR at of 1200 pmol/m'/s (Table 1). Amux values were significantly different from each
other (one-way ANOVA, P = 0.0296), while the PARs at the A,nt,x values were not significantly different
between treatments (one-way ANOVA, P = 0.2861).
The quantum yield efficiency or initial slope or IS) for shade leaves (0.030 ± 0.010 pmol
C02/( pmol quanta) was not significantly different from that of sun leaves (0.032 ±0.010 pmol C02/(pmol
quanta) (one-way ANOVA, P = 0.0677, Table 1). The liglit compensation point (Lcp), the light saturation
point (Lsp) and dai*k respiration (Rj) for shade leaves were not significantly different from the sun leaves
(one-way ANOVA, P = 0. 1431, 0.2618 and 0.0758 respectively. Table 1).
There were no overall significant differences in mean transpiration rate between sun and shade
leaves (repeated-measures ANOVA; P = 0.2274) (Table 1). However, the transpiration rate for sun
leaves increased from 0.34 ±0.12 mmol H20/m"/s at the lowest light level to 1.47 ± 0.12 mmol H20/nr/s
at the highest light level tested with a few significant differences. Usually significant differences were
between the lowest light level and the highest (one-way ANOVA; P < 0.000 1 ; Tukey - Kramer HSD; P <
0.05) (data not shown). The mean transpiration rate for shade leaves increased from 0.36 ± 0.08 mmol
H20/m“/s to 1.04 ± 0.08 mmol H20/mVs with a similar trend in significant differences (one-way
ANOVA; P < 0.0001; Tukey - Kramer HSD; P < 0.05) (data not shown).
There were no overall significant differences in mean stomatal conductance between sun and
shade leaves (repeated-measures ANOVA; P = 0.9305) (Table 1). However, the conductance for sun
leaves mcreased from 0.01 ± 0.01 mol H20/nr/s to 0.05 ± 0.01 mol HoO/m^/s with few significant
differences. Usually significant differences were between the lowest light level and the highest (one-way
ANOVA; P < 0.0001; Tukey - Kramer HSD; P < 0.05) (data not shown). The conductance for shade
leaves increased from 0.02 ± 0.05 mol H20/m~/s to 0.04 ± 0.05 mol H20/m"/s with few significant
differences and trends similar to transpiration (one-way ANOVA; P < 0.0001; Tukey - Kramer HSD; P <
0.01) (data not shown).
DISCUSSION
Acer grandidentatwn seedlings grew best m the 40 % light treatment, which was 705 ± 22
pmol/m/s, and most closely matches the light levels found below an A. grandidentatum canopy at Lost
Maples State Natural Area. This was where there were higher numbers of A. grandidentatum saplings and
mature trees and suggests better seedling survival (Nelson Dickerson and Van Auken 2016). All of the
plants survived in the 40 % light treatment, ^^frereas none of the A. grandidentatwn seedlings in the
highest light exposure survived, supporting the hypothesis tliat this species is a shade and not a sun
species. Light levels far above a plant’s light saturation point that do not cause an increase in
Phytologia (Jan 19, 2017) 99(1)
15
photosynthesis, may cause damage to leaf tissues and the photosynthetic apparatus or increase water loss
to the pomt of wilting and possibly mortality (Crawley 1997).
Table 1. Mean ± one standard error for the maximum net photosynthetic rates light level (PAR) at
the light saturation (Lsa,), light compensation points {Lcp), dark respiration rates (R^/), initial slope or
quantimi yield efficiency (IS\ mean stomatal conductance (^ 5 ) and mean transpiration rate (E) of A.
grandidentatiini leaves foimd m full sun and shade. Stars mdicate a significant difference between values
for the two treatments (one-way ANOVA; P < 0.05).
Parameter
Shade leaves
Sun leaves
ATOaTpmol CO 2 / m/s)
3.89 ±0.36*
5.23 ±0.36*
PAR at A,,k,.v (pmol/m/s)
880 ±198
1200 ±198
Mean photosynthetic rate
2.27 ±0.23
2.94 ±0.23
Lsat ( pmol/m/s)
139.11 ± 18.48
1 70.67 ± 18.48
Lcp{ pmol/m/s )
9.74± 2.22
14.83±2.22
Re! (pmol C 02 /m"s)
0.32 ± 0.05
0.47 ± 0.05
IS (pmol C02/(pmol quanta)
0.032 ±0.010
0.030 ±0.010
gs (mol H 20 /m/s)
0.04 ±0.01
0.04 ±0.01
E (mmol H 2 0/m/s)
0.83 ±0.08
0.99 ±0.12
When plants are grown with msufficient light, they may have decreased leaf area, basal diameter
and dry mass (Jones and McLeod 1989), but may increase their shoot height or decrease their leaf to
shoot ratios (Holt 1995). Acer grand identatuin seedlings in the present study that received 20 % of
ambient sunlight had high mortality and low growth (Figure 2). Similar deep shading of A. saccharum
and Aesculus glabra seedlings in early spring led to 80 % mortality after tliree years, compared to 27 %
mortality in control plants (Augspurger 2008). Though A. grandidentatum seems to be best characterized
as a shade plant, it still requires approximate 40 % sunlight to compensate for its respiration to allow
growth and surs/ival. The majority of growth for all experimental seedlings in the current study occurred
in spring, during the first half of the experiment (not presented ). Growth for all plants slow ed through the
intense heat and light of summer, and most mortality' was observed late m the experiment (Nelson
Dickerson 2011).
The current results differ somewhat from similar studies done on the closely related A.
saccharum. First-year A. saccharum seedlings grown at 13, 25, 45 and 100 % sunliglit for one year
increased their number of leaves and dry mass as light level increased, with maximum values found at
100 % sunlight (Logan and Krotkov 1968). The study was conducted in Ontai'io, Canada, and does not
disclose the actual light levels used in the experiment. It is quite possible that the amount of sunlight
received by these plants was lower than tlie 100 % suggested. In addition, ambient temperature was
significantly different tlian used in the current experiment due to the higher latitude and shorter day length
or possibly other factors that were not the same as in the present study. However, A. saccharum is
considered to be a shade plant (Logan and Krotkov 1968; Ellsworth et al. 1994; Kwit et al. 2010).
Gas exchange rates of members of the genus Acer indicates the genus includes both shade
tolerant and shade intolerant species (Morrison and Mauck 2007; Verdu and Climent 2007). While no
infonnation on the photosynthetic parameters of A. grandidentatum have been identified in the literature,
it is assumed to be at least moderately shade tolerant because of its distribution in protected canyon
bottoms and as an understor>' late succession species (Conell and Johnston 1970; Bazzaz and Carlson
1982; Nelson Dickerson and Van Auken 2016). The slow growth of A. grandidentatum also suggests
shade tolerance, as photosynthetic parameters are closely tied to relative growth rates (Coley et al. 1985;
Poorter 1990; Tollefson 2()06; Van Auken et al. 2016).
16
Phytologia (Jan 19, 2017) 99(1)
Acer grandidentatum maximum photosynthetic rates of 3.89 ± 0.36 pmol COi/m /s for shade
leaves and 5.23 ± 0.36 pmol C02/nr/s reported for sun leaves in the present study are consistent with
classification of as a shade plant. Acer grandidentatum sun plants found at higher elevations, higher
rainfall and lower temperatures appear to have higher values in fiill sun (Van Auken and Bush, in
preparation, unpublished), but plants in shade had similar low A,„av values. Succession in many cases is
driven by temporal differences in resource availability and paiticulai'ly by changes in available nutrients,
especially nitrogen, and light levels (Tilman 1985; Van Auken and Bush 2013).
Early succession sites usually have high light levels and low soil nitrogen. Usually early
successional species ai*e shade intolerant and late successional species are shade tolerant (Boai'dman
1977; Tilman 1985; Mooney and Ehleringer 1997; Valladaies and Niinements 2008; Van Auken and
Bush 2013). As successional time passes and communities mature, increased canopy shading decreases
available light at the soil surface and shade tolerant and higher soil nitrogen requiring plants become more
common (Tilman 1985; Bush and Van Auken 1986; Van Auken and Bush 2013). Early successional
species exhibit higher rates of photosynthesis, transpiration, and conductance than late successional
species, while late successional or climax community species are more likely to be shade tolerant and
reach tlieir light saturation points at much lower light levels (Horn 1974; Fumya and Van Auken 2009,
2010; Wayne and Van Auken 2009; Van Auken and Bush 2011). Early succession sites also have greater
variability in abiotic conditions, such as swinging between environmental extremes, so early successional
plants frequently have greater plasticity in their adaptive responses than late successional species (Horn
1974; Bazzaz and Carlson 1982; Holt 1995; Hull 2002; Van Auken and Bush 201 1).
Bazzaz and Carlson (1982) measured photosynthetic rates of flill sun and shade leaves of fourteen
speeies. They found that the difference between initial slope, light compensation point, and dark
respiration for foil sun and shade leaves was much greater for herbaceous early succession species than
for late succession hardwood species. The values they reported for late successional hardwoods are
similar to foe values obtained for A. grandidentatum in the present study (Table 1). The present .study
found no significant differences between most variables for sun and shade leaves, eonsistent with
observations that A. grandidentatum is a late sueeession speeies (Bazzaz and Carlson 1982; Hull 2002).
Light response curves for A. sacchamm seedlings in clearings had A^ax values of 3.32 pmol
C 02 /mVs, w'hile understory individuals had values of 1.81 pmol C 02 /m“/s, the only factors that were
significantly different between locations (Ellsworth and Reich 1992). We found similar values for A.
grandidentauun m the present study. These tv\o species may be able to increase their photosynthetic rate
to talce advantage of sunflecks, short term increases in light availability, but limited data is available.
Acer grandidentatum seems to exhibit lower plasticity in photosynthetic rate than other woody species,
which may affect its growth as well as its ability to become a dominant member of the canopy (Hull
2002). Photosynthetic response curves below a forest canopy measured at four different times in a single
growing season found that most light response parameters decreased during foe growing season and
affinned that A, saccharum is shade tolerant, and seedlings must survive most of foeir first year in a
densely shaded forest eanopy (Kwit et al. 2010).
Measured transpiration rates {E) and stomatal eonduetanee (gs) rates for A. grandidentatum were
low' compared to values for shade intolerant species (Boardman 1977; Bsoul et al. 2007). These rates
indicated the stomates were open and CO 2 uptake was probably normal. Rates are consistent with values
for shade tolerant species but not shade intolerant species (Horn 1974; Boaifonan 1977; Bazzaz and
Carlson 1982; Tilman 1985; Holt 1995; Mooney and Ehleringer 1997; Valladares and Niinements 2008;
Hull 2002; Van Auken and Bush 2013).
Phytologia (Jan 19, 2017) 99(1)
17
Photosynthetic rates of A. grandidentatum were lower at high light levels than those of most other
dominant plant species in the community at Lost Maples State Natural Area, but not at low light sub-
canopy conditions (Furuya 2007; Grunstra 2011; Grunstra and Van Auken 2015). Differing light
requirements have been shown to affect succession and plant community composition (Bush and Van
Auken 1986; Wayne and Van Auken 2009; Van Auken and Bush 2011). Plants with low photosynthetic
rates may experience difficulty growing below the canopy and then through the canopy without a
disturbance to canopy plants. When this lowered potential is combined with browsing pressure, the effect
can become even stronger (Van Auken and Bush 2009). Community composition in Sabinal Canyon in
Lost Maples State Natural Area and other central Texas communities are likely affected by complex
interaction of inherent photosynthetic capacities and abiotic requirements of the species present. This
would including preferential feeding of large herbivores, and the effect of that herbivory on the biotic and
abiotic conditions present in the environment. There are a number of species found in these central Texas
communities that can grow at high light levels, but most cannot grow in deep shade below a closed
canopy (Furuya 2007; Grunstra 2011; Grunstra and Van Auken 2015).
THE FUTURE
Populations of A. grandidentatum in central Texas are relatively rare and are really outlier
populations. Management of these populations in the past has mostly been hap-hazard at best and
dependent on the whims of owners of properties where they have been found. Understanding that they are
understory/sub-canopy species or shade species was unknown until the present study. Sensitivity to native
and domestic herbivores has been suspected for many years but not demonstrated until very recently.
What will happen to these populations in the future? This is uncertain and difficult to predict. If their
reproductive cycle is continually disrupted, they will become extinct in central Texas. If herbivory by
native and domestic species is not reduced the same thing will happen. What is the timeline of the
potential extinction of these isolated native populations? This is uncertain at this time. It is hard to say
because apparently individuals can live for hundreds of years and the death rate of adults is unknown and
the rate of recruitment of juveniles into these populations is not known either.
ACKNOWLEDGEMENTS
We thank Jeffrey Jackson, Vonnie Jackson, Matthew Grunstra and Anne Adams for reading an
earlier draft of this manuscript and making many helpful corrections and suggestions. We also thank
Amy Moulton and Jacque Keller for help formatting this manuscript.
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Phytologia (Jan 19, 2017) 99(1)
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U)
w
(C
E
a
20% 40% 60% 100%
Light Level (% of Ambient)
Figure 1. Mean aboveground (■), belowground and total dry mass of Acer grandidentatum at
varying light levels as a pereentage of ambient sunlight. Bars indieate ± one standard deviation of the
mean. Different letters on the same line indieate signifieant differenees for that faetor (one-way ANOVA;
P < 0.05, Tukey - Kramer HSD; P < 0.05).
1 *
0.9 -
^ 0.8 -
I' 0.7 -
t o.e -
I 0.5 -
0.4 -
0.3 -
0.2 -
0.1 -
0 -
20% 40% 60% 100%
Light Level (% of Ambient)
Figure 2. Relative mortality (1=100%) of Acer grandidentatum seedlings grown at varying light levels is
presented as a pereentage of ambient sunlight. The * indieates values whieh were signifieantly different
from the mean (ANOM for Proportions, a = 0.05, LDL [lower deteetion limit] = 0, UDL [upper deteetion
limit] = 0.868). There were no mortalities in the 40% light treatment.
Phytologia (Jan 19, 2017) 99(1)
21
Figure 3. A. Mean photo synthetie rate of full sun and shaded leaves of A. grandidentatum as a funetion
of light level (PAR). B. The lower portion of graph A between 0 and 100 PAR. Upperease letters
represent values for sun leaves, while lowerease letters represent values for shaded leaves. Different
upper or lowerease letters indieate signifieant differenees within the eurve (one-way ANOVA; P < 0.0001;
Tukey -Kramer HSD; P < 0.05). Error bars represent ± one standard error.
22
Phytologia (Jan 19, 2017) 99(1)
Discovery of Juniperus sabina var. balkanensis R. P. Adams and A. N. Tashev
in western Turkey (Anatolia)
Robert P. Adams
Biology Department, Baylor University, Gmver Lab, Gmver, TX 79040, USA
Robert Adanis@bavlor.edu
Adam Boratynski
Institute of Dendrology, Polish Aeademy of Seienees, Parkowa 5, 52-035, Komik, Poland
Tugrul Mataraci
Eskidji Mtiz. A§., Sanayi Cad. Vadi SokakNo:2, Yenibosna-Bah 9 elievler, Istanbul, Turkey
Alexander N. Tashev
University of Forestry, Dept, of Dendrology, 10, Kliment Oehridsky Blvd., 1756 Sofia, Bulgaria
and
Andrea E. Schwarzbach
Department of Health and Biomedieal Seienees, University of Texas - Rio Grande Valley,
Brownsville, TX 78520, USA.
ABSTRACT
Additional analyses of tmS-tmG and mDNA from herbarium speeimens from Europe revealed
the presenee of J. sabina var. balkanensis in western Turkey near Izmir and expands the range previously
known only from Bulgaria and adjaeent mountains in Greeee. A more detailed map of the taxon's
distribution is presented. Published on-line www.phytologia.org Phytologia 99(1): 22-21 (Jan 19, 2017).
ISSN 030319430.
KEY WORDS: Juniperus sabina var. balkanensis, J. sabina, distribution, nrDNA, tmS-tmG, ehloroplast
eapture.
Reeently, Adams et al. (2016) reported on the eapture of J. thurifera (or an aneestor) ehloroplast
by J. sabina var. balkanensis. Chloroplast eapture has been rarely reported in eonifers. In Pinus and
other eonifers, Hipkins et al. (1994) eoneluded that "past hybridization and assoeiated 'ehloroplast
eapture' ean eonfiise the phytogenies of eonifers." Bouille et al. (2011) found signifieant topologieal
differenees in phylogenetie trees based on epDNA (vs. mtDNA sequenees) in Picea that suggested
organelle eapture.
In Juniperus, Terry et al. (2000) suggested that ehloroplast eapture was involved in the
distribution of ep haplotypes in J. osteosperma in western North Ameriea. More reeently, Adams (2015 a,
b) found widespread hybridization and introgression between J. maritima and J. scopulorum in the
Paeifie northwest, with introgression from J. maritima into J. scopulorum eastward into Montana. The
disparity between epDNA and nuelear markers (nrDNA and maldehy) suggested that ep eapture had
oeeurred.
The Juniperus of seetion Sabina, of the eastern hemisphere, ean be divided into two groups based
on the number of seeds per female eone (often ealled berries) and female eone shape. The single
seed/eone (single-seeded) Juniperus of the eastern hemisphere have eones that are ovoid with a
notieeable pointed tip, whereas the multi-seeded Juniperus are generally globose and often have an
Phytologia (Jan 19, 2017) 99(1)
23
irregular surface (Adams 2014). Juniperus sabina L. is a smooth leaf-margined, multi-seeded juniper of
the eastern hemisphere. It is very widely distributed from Spain through Europe to Kazakhstan, western
China, Mongolia and Siberia (Fig. 1). Juniperus sabina has a range that is discontinuous between Europe
and central Asia; the species is generally a shrub less than 1 m tall and ranges up to 1-2 m wide. But in
the Sierra Nevada of Spain, it forms a horizontal shrub.
Fig. 1. Distribution (shaded areas) of J. sabina. x = outlying populations of J. sabina.
Adams et al. (2016) showed that mDNA (ITS) did not resolve J. sabina populations due to the
lack of sequence variation.
However, their analyses (Adams et al., 2016) of cp DNA (petN-psbM, tmSG, tmDt, tmEF)
revealed that J. sabina contained two kinds of cpDNA: typical J. sabina and J. sabina var. balkanensis
cpDNA in a clade with J. thurifera (Fig. 2).
It might be noted that J. sabina from Kazakhstan and Xinjiang form a clade (Fig. 2). The use of
four cp regions resulted in a clade of the junipers from the western hemisphere (box. Fig. 2).
In order to investigate the amount of divergence of the 'balkanensis' chloroplast from that of
present day J. thurifera, a minimum spanning network was computed using both SNPs and indels, herein
called mutations. This analysis found 52 mutations within the set: J. sabina {sensu stricto), J. sabina J. s.
var. balkanensis and J. thurifera. The minimum spanning network (Fig. 3) shows that all the
'balkanensis' plants differ by only 6-8 mutations from J. thurifera chloroplast. However, the nearest link
connecting 'balkanensis' to J. sabina {sensu stricto) is 36 mutations!
Notice (Fig. 3) that Azerbaijan/ Mongolia accessions group with Kazakhstan/ Xinjiang and this
group differs by 7 mutations from the Europe/ Algeria group. This suggests that J. sabina in central Asia
may be a different variety of J. sabina. That needs to be examined in more detail (in progress).
24
Phytologia (Jan 19, 2017) 99(1)
_^7846 communis - outgroup
7847 communis
88
100 '
53
100
oo ■
\ 100 .
8785 excelsa
100
— 6184 pracera
“ 8761 polycarpos
8757 turcomanica
8224 seravschanica, Kaz.
8483 seravschanica, Pak.
'5645 foetidissima
8688 chin v sargentii
8535 chinensis
.8683. chin_v.p,rDc.u[tibjet)&.
13726 'balkanensis' Bulg.
13726 'balkanensis' Bulg.
thur. v aWcana
7083 thurifera
97
100
66
14722 ’balkanensis' Buig.
94*" 1 4728 ’balkan ensis ’ Greece
IQO I 7077 phoenicea
' 7202 turbinata
9061 tsukusiensis V taiwanensis
1 “ 7820 semiglobosa v jarkendensis
100 * — 8210 semiglobosa
100 I 13633 microsperma
100
8532 erectopatens
89
■11056 maritima
— 10231 virginiana
93
100
— 10895 scopulorum
10247 blancoi v huehuentensis
88
95
98
western
hemi,
junipers
66849 blancoi
8701 blancoi v mucronata
35 r gradlior v ekmanii
— U 7656 gracilior v urbaniana
^5284 gracilior v saxtcola
— 11080 bermudiana
- 7664 gracilior
— 9186 virginiana v. silidcola
“ 7096 horizontalis
56
99
5358 barbadensis
5281 barb, v lucavana
76
100
Bayesian
petN-psbM,
trnDT, trnLF
3114 bp
tree
trnSG,
8806 tsukusiensis
- 7254 davurica v mongolensis
10347 davurica v arenaria
10348 davurica v arenaria
-7252 davurica
7253 davurica
- 72_55 da_vurica v mongolensis
r*7811 sabina. Kazakhstan
^ "I 7812 sabina, Kazakhstan
1 7836 sabina, Xingiang
* 7836 sabina, Xingiang
“ 7587 sabina, Altai Mtn, Mong.
7585 sabina, Altai Mtn Mong.
7586 sabina, Altai Mtn. Mong.
7197 sabina. Sierra Nevada, Spain
100 r ^^99 sabina, Sierra Nevada, Spain
7573 sabina, Pyrenees
7574 sabina, Pyrenees
7612 sabina, Switzerland
7614 sabina, Switzerland
14316 sabina, Azerbaijan
14317 sabina, Azerbaijan
98
Figure 2. Bayesian analysis based on four ep regions (adapted from Adams et af, 2016).
Adams et al. (2016) eoneluded that J. sabina var. balkanensis eaptured the ehloroplast of an
aneestor of the thurifera lineage during an aneient hybridization event at a time when speeies
distributions overlapped. Beeause var. balkanensis has morphology almost identieal to J. sabina {sensu
stricto), this hybridization event was likely followed by sueeessive baekerosses to J. sabina after the
Phytologia (Jan 19, 2017) 99(1)
25
hybridization event, resulting in a nuelear genome, ineluding morphology, that is nearly identieal to J.
sabina (sensu stricto). In faet, Adams et al, 2016 found in the mDNA analysis that J. s. var balkanensis
was elearly interspersed in a elade with other J. sabina. So it is not surprising that a eomparison of the
morphology of J. sabina and J. s. var. balkanensis, has, to date, revealed only a few quantitative
differenees (Adams et al. 2016, Table 1).
Figure 3. Minimum spanning network based on 52 mutations (SNPs + indels) in 4 ep markers (3 1 14 bp).
The numbers next to the lines are the number of mutations for that link. The dotted line eonneets the
thurifera ep taxa to the sabina ep taxa by 36 mutations. The dashed line is the seeond nearest neighbor of
J. sabina to J. davurica ep type. (8 mutations).
Juniperus sabina var. balkanensis is known only from sloping roeky limestone, at 1240 - 1630m,
in the mountains of Bulgaria and northern Greeee (Fig. 4). Adams et al. (2016) postulated that it may
oeeur northward into Romania, westward into Maeedonia and/ or eastward into northern Turkey.
26
Phytologia (Jan 19, 2017) 99(1)
The purpose of the present paper is to report on a broader sampling of J. sabina from herbarium
speeimens to more preeisely determine the distribution of J. sabina var. balkanensis.
Fig. 4. Habit and habitat of J.
var. balkanensis in the eastern
Rhodopes mountains, Bulgaria.
Juniperus communis, eolumnar
trees, are in the baekground.
MATERIAL AND METHODS
Speeimens used in this study (speeies, popn. id., loeation, eolleetion numbers): J. chinensis, CH,
Lanzhou, Gansu, China, Adams 6765-6767', J. sabina'. (SN), Sierra Nevada, Spain, Adams 7197, 7199,
7200', (PY), Pyrenees Mtns., Spain/ Franee border, Adams 7573-7577', (SW), Switzerland, Adams 7611,
7612, 7614, 7615', TS, Tian Shan Mtns., Xinjiang, China, Adams 7836-7838', Mongolia, Altai Mtns.,
Adams 7585-7587', Kazakhstan, PmAfior, Adams 7811-7812', Azerbaijan: Adams 14316-14320',
J. davurica (DV), 15 km se Ulan Bator, Mongolia, Adams 7252, 7253, 7601', J. davurica var. arenaria
(AR) sand dunes. Lake Qinghai, Qinghai, China, Adams 10347-10352', river bank, Gansu, J-Q. Liu and
Adams 10354-10356', J. davurica var. mongolensis (MS) sand dunes, 80 km sw Ulan Bator, Mongolia,
Adams 7254-7256',
Collections of tcocon with non-J. sabina cpDNA in Adams, Schwarzbach and Tashev (2016): (acronyms
used in Fig. 7)
Bulgaria and Greece
B1-B5 Eastern Rhodopes, Bulgaria, Adams 13725-13729 (A. Tashev 2012-1-5)',
B6 Central Stara Plania, Sokolna reserve, Bulgaria, Adams 14721 (A. Tashev 2015 Balkan 7;
B7-B9, Ba, Bb Rila Mountain, Bulgaria, Adams 14722-14726 (A. Tashev 2015 Rila 1.1-1. 3, 2. 1-2.2)',
G1-G5 Mt. Tsena, Gxqqcq, Adams 14727-14731 {A. Tashev 2015 So. 1-5 Tsena)',
Samples new for this study: (with Lab Acc. ID = Adams xxxxx)
Austria
14872 Austria, Alps, Otztal, Zwiselstein, N 46.935°, El 1.039°, 1650-1700m alt., leg. K. Boratyhska,
A.Boratyhski, 2015, 15.001, KOR 51592, female
14873 Austria, Alps, Otztal, Below Solden, N 46.994°, El 1.012°, 1300 alt., leg. K. Boratyhska,
A.Boratyhski, 2015, 15.005, KOR 51596, male
14874 Austria, Alps, Otztal, Below Solden, N 46.994°, El 1.012°, 1300 alt., leg. K. Boratyhska,
A.Boratyhski, 2015, 15.005, KOR 51595, female
France
14863 Franee, Alps de Dauphine, St. Crepin, N 44.71°, E 6.61°, ea 1000m alt, leg. A. Boratyhski, K.
Boratyhska 2003, 03.19.116, KOR 43778, female
Phytologia (Jan 19, 2017) 99(1)
27
Italy
14870 Italy, Alps, Val d’Aosta, Introd, Les Combes, N 45,689°, E 7.166°, 1250 m alt. Lag. K.
Boratynska, A. Boratynski, 15.014. KOR 51590, female
14871 Italy, Alps, Val d’Aosta. Introd, Les Combes, N 45.689°, E 7.166°, 1250 m alt. Lag. K.
Boratynska, A. Boratynski, 15.013. KOR 51589, male
Poland
14858 Poland, Carpathians, Pieniny National Park, Facimiech, N 49.40°, E 20.43°, ca 600m alt. From
specimen propagated vegetatively about 2005 and planted in dendrological garden of Forest Botany
Chair, Forest Faculty, Poznan University of Lite Sciences
Russia
14865 Russia, Altay, Aktru Valley, SWW of Bielucha Mt., ca. N 49.80°, E 86.40°, 2500m alt., leg.
FalNnowicz W., 2010. KOR 4796, female.
Spain
14860 Spain, Cuenca, Serrana de Cuenca, between Tragacete and La Cueva (Vega de Cordorno), N
40.433°, W 1.905°, ca 1450 m alt, Ig. Boratynska K., Boratynski A., 2006, SP.06.026, KOR 44733,
female
14862 Spain, Teruel, Puerto de Cabigordo near Cedrillas E of Teruel, N 40.41°, WO. 95°, ca 1500m alt.,
Leg. A. Borat^Tiski, K. Boratynska, hlS_01.03.17, KOR 43212,
14864 Spain, Sierra Nevada, Veleta Mt., above Alberque Universitario, N 37.09°, W 3.38°, ca 2500m alt.,
leg. A. Boratynski 1991, KOR 25299
14866 Spain, Sierra Nevada, Monte Ahi de Cara, N 37.13°, W3.43°, 1900-2000m alt., leg. A.Boratyhski,
Ja. Didukh., D.Tomaszewski, Z. Boratynski, KOR 46220, female
14869 Spain, Leon, Los Banos de Luna, N 42.88°, W 5.87°, 1 150- 1200m alt., leg. K.Boratyhska, A.
Boratynski, 2015, KOR 51542, female
14875 Spain, Sierra de Albanacin, S of Brochales, N 40.50°, W 1.57°, ca 1600m alt., leg. A. Boratynski,
K. Boratynska, 2006. female
14876 Spain, Aragon, Moncayo, N 41.77°, W 1.80°, ca 1900-2000m alt, leg. D. Gomez, 2004, female
Switzerland
14867 Switzerland, Alps, Visp, Aussenberg, N 46.31°, E 7.87°, ca 950-1000m alt., leg. K.Boratyhska,
A.Boratyhski, 2015, 15.016, KOR 51570, female
14868 Switzerland, Alps, Visp, Aussenberg, N 46.31°, E 7.87°, ca 950-1000m alt., leg. K.Boratyhska,
A.Boratyhski, 2015, 15.017, KOR 51581, male
Turkey
14861 Turkey, Manisa. Spil Dagi Milli Parki (National Park) (Tas Suret), N38.55°, L 27.42°, ca 1250 m
alt., leg. A. Boralyhski, K. Boratyhska, 2005, TU_05/55, KOR 44573, female
14934 Tiukey, Manisa, Spil Dagi Milli Pai'ki (National Park), N38°, 57', E 27° 41', 1024 m., Tii^‘ul
Mataraci 2016-1
14928 Turkey, Gumushane, Kurtun, Alctas village, Karakaya (Northeast Anatolia), 40° 36' 03" N, 38° 53'
21" E., 2376 m. Coll. A. Kandemir 10745.
Ukraine
14859 Ulsiaine, Crimea, Chatyr Dag, N 44.773°, E 34.313°, 1100-1200m alt. Eg. A. Boratyhsld, G.
Iszkulo, A. Lewandowski, 2006. UA06.007, KOR 45572
Voucher specimens for all collections are deposited at Baylor University Herbarium (BAYLU)
and Herbarium (University of Forestry, Sofia, Bulgaria).
One gram (fresh weight) of the foliage was placed in 20 g of activated silica gel and transported
to the lab, thence stored at -20" C until the DNA was extracted. DNA was extracted from juniper leaves
by use of a Qiagen mini-plant kit (Qiagen, Valencia, CA) as per manufacturer's instructions.
Amplifications were perfonned in 30 pi reactions using 6 ng of genomic DNA, 1.5 units Epi-Centre Fail-
Safe Taq polymerase, 15 pi 2x buffer E (petN, tniD-T, trriL-F, tmS-G) or K (iirDNA) (final
concentration: 50 inM KCl, 50 inM Tris-HCl (pH 8.3), 200 pM each dNTP, plus Epi-Centre proprietary
28
Phytologia (Jan 19, 2017) 99(1)
enhancers with 1.5 - 3.5 mM MgCl 2 according to the buffer used) 1.8 gM each primer. See Adams,
Bartel and Price (2009) for the ITS and petN-psbM primers utilized. The primers for tmD-tmT, tmL-tmF
and tmS-tmG regions have been previously reported (Adams and Kauffmann, 2010). The PCR reaction
was subjected to purification by agarose gel electrophoresis. In each case, the band was excised and
purified using a Qiagen QlAquick gel extraction kit (Qiagen, Valencia, CA). The gel purified DNA band
with the appropriate sequencing primer was sent to McLab Inc. (San Francisco) for sequencing.
Sequences for both strands were edited and a consensus sequence was produced using Chromas, version
2.31 (Technelysium Pty Ltd.) or Sequencher v. 5 (genecodes.com). Sequence datasets were analyzed
using Geneious v. R7 (Biomatters. Available from http://www.geneious.com/) . the MAFFT alignment
program. Further analyses utilized the Bayesian analysis software Mr. Bayes v.3.1 (Ronquist and
Huelsenbeck 2003). For phylogenetic analyses, appropriate nucleotide substitution models were selected
using Modeltest v3.7 (Posada and Crandall 1998) and Akaike's infomiation criterion. Minimum spanning
networks were constructed from mutational events (ME) data using PCODNA software (Adams, Bartel
and Price, 2009; Adams, 1975; Veldman, 1967).
RESULTS
The results of this study (and the previous, Adams et al. ,2016 study) are given in Table 1. The
distribution of J. sabina var. balkanensis and J. sabina is shown in Fig. 5. The distribution of J. thurifera
O balkanensis i
thurifera
chloroplast
Figure 5. Distribution of J. sabina var. balkanensis and typical J. sabina chloroplast. The present day
distributions of J. thurifera and var. africana (in north Africa) are shown in the insert on the lower left.
Phytologia (Jan 19, 2017) 99(1)
29
is presented in the insert, lower left (Fig. 5). It appears that J. s. var. balkanensis has a quite restricted
range. Additional samples are needed from Romania, Turkey and northwesterly from Albania/
Macedonia northwesterly to Slovenia to determine the distribution more precisely.
At present level of understanding, the distributions of J. s. var. balkanensis and J. thuhfera do not
appear to overlap, negating modem hybridization. However, there were large changes in plant
distributions in the Pleistocene and earlier, it seem probable that J. thurifera-\\k& ancestors were
sympatric with J. sabina, and presenting opportunities for cliloroplast capture from J. thurifera.
ACKNOWLEDGEMENTS
Thanks of A. Kandemir for the specimen of J. sabina from Aktas village, northern Turkey. This
research was supported with ftmds provided by Baylor University.
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geographic speciation and reticulation as driving factors for the diversification of the genus Picea.
Tree Genetics & Genomes 7: 469-484.
Hipkins, V. D., K. V. Knitovskii and S. H. Strauss. 1994. Organelle genomes in conifers; stmcture,
evolution, and diversity. Forest Genetics 1: 179-189.
Posada, D. andK. A. Crandall. 1998. MODELTEST: testing the model of DNA substitution.
Bioinfonnatics 14; 817-818.
Ronquist. F. and J. P. Huelsenbeck. 2003. Mi'Bayes 3: Bayesian phylogenetic inference under mixed
models, Bioinfonnatics 19: 1572-1574.
Terr}', R. C., R. S. Nowak and R. J. Tausch. 2000. Genetic variation in chloroplast and nuclear
ribosomal DNA in Utah juniper {Jimiperus osteosperma, Cupressaceae): evidence of mterspecific
gene flow. Am. J. Bot. 87: 250-258.
Veldman, D. J., 1967. Fortran programming for the behavioral sciences. Holt, Rinehart and Winston
Publ, NY.
30
Phytologia (Jati 19, 2017) 99(1)
Table 1. Classification of J. sabina specimens based on tmS-tmG (plus petN-psbM, tmDT, tmLF) and
nrDNA (ITS).
Lab Acc. #, Location
trnSG (cp genome)
classification
nrDNA
classification
13725 Bulgaria, eastern Rhodopes
V. balkanensis
V. sabina
13726 Bulgaria, eastern Rhodopes
V. balkanensis
V. sabina
13727 Bulgaria, eastern Rhodopes
V. balkanensis
V. sabina
13728 Bulgana, eastern Rhodopes
V. balkanensis
V. sabina
13729 Bulgaria, eastern Rhodopes
V. balkanensis
V. sabina
14721 Bulgaria, Sokolna reser\-e
V. balkanensis
V. sabina
14722 Bulgaria, Rila Mtn.
V. balkanensis
V. sabina
14723 Bulgaria, Rila Mtn.
V. balkanensis
V. sabina
14724 Bulgaria, Rila Mtn.
V. balkanensis
V. sabina
14725 Bulgaria, Rila Mtn.
V. balkanensis
V. sabina
14726 Bulgaria, Rila Mtn.
V. balkanensis
V. sabina
14727 Greece, Tsena Mt.
V. balkanensis
V. sabina
14728 Greece, Tsena Mt.
V. balkanensis
V. sabina
14729 Greece, Tsena Mt.
V. balkanensis
V. sabina
14730 Greece, Tsena Mt.
V. balkanensis
V. sabina
14731 Greece., Tsena Mt.
V. balkanensis
V. sabina
14934 w Turkey, Spil Dagi Milli Park!
V. balkanensis
V. sabina
14861 w Turkey, Spil Dagi Milli Parki
V. balkanensis
V. sabina
13167 Algeria
V. sabina
V. sabina
13168 Algeria
V. sabina
V. sabina
14872 Austria, Otztal. Zwiselstein
V. sabina
V. sabina
14873 Austria, Otztal, Below Sdlden,
V. sabina
V. sabina
14874 Austria, Otztal, Below Sdlden,
V. sabina
V. sabina
14316 Azerbaijan
V. sabina
V. sabina
14317 Azerbaijan
V. sabina
V. sabina
7836 China. Heaven Lake, Xinjiang
V. sabina
V. sabina
7837 China, Heaven Lake, Xinjiang
V. sabina
V. sabina
14863 France, Alps de Dauphine
V. sabina
V. sabina
7573 France, P>Tennes Mtns
V. sabina
V. sabina
7574 France, lA rennes Mtns
V. sabina
V. sabina
14870 Italy, Val d’Aosta, Alps
V. sabina
V. sabina
14871 Italy, Val d' Aosta, Alps
V. sabina
V. sabina
7811 Kazakhstan, Paniflor
V. sabina
V. sabina
7812 Kazakhstan, Paniflor
V. sabina
V. sabina
7585 Monsolia, Altair Mtns
V. sabina
V. sabina
7586 Moneolia, Altair Mtns
V. sabina
V. sabina
7587 Monad ia. Altair Mtns
V. sabina
v. sabina
14858 Poland, Pieniny N.P.,
V. sabina
V. sabina
14865 Russia. Altav Mtn.
V. sabina
V. sabina
7197 Spain, Sierra Nevada
V, sabina
V. sabina
7199 Spain, Sierra Ne^'ada
V. sabina
V. sabina
14860 Spain. Serrana de Cuenca
V. sabina
V. sabina
14862 Spain, Teruel
V. sabina
V. sabina
14864 Spain, Sierra Nevada
V. sabina
V. sabina
14866 Spain, Sierra Nevada
V. sabina
V. sabina
14869 Spam. Los Barios de Luna
V. sabina
V. sabina
14875 Spam, Sierra de Albarracin
V. sabina
V. sabina
14876 Spain, Aragon, Moncayo
V. sabina
V. sabina
Phytologia (Jan 19, 2017) 99(1)
31
761 1 Switzerland, Alps
V. sabina
V. sabina
7612 Switzerland, Alps
V. sabina
V. sabina
7614 Switzerland , Alps
V. sabina
V. sabina
14867 Switzerland, Aussenberg
V. sabina
V. sabina
14868 Switzerland, Aussenberg
V. sabina
V. sabina
14938 northeast Turkey
V. sabina
V. sabina
14859 Ukraine, Crimea, Chatry Dag
V. sabina
V. sabina
32
Phytologia (Jan 19, 2017) 99(1)
The effects of different concentrations of gibberellic acid (GA3) on seed germination of
Helianthus annuus and H. petiolaris
Robert P. Adams and Amy K. TeBeest
Biology Department, Gruver Lab, Baylor University, Gruver, TX 79040, USA
robert_adams@baylor.edu
ABSTRACT
Germination tests were conducted by soaking native, wild Helianthus annuus and H. petiolaris
seeds in various concentrations of gibberellic acid (GAS) for 1 week, 4°C. For H. annuus, the most
effective concentrations of GA3 were 1000 ppm (61.7%) and 500 ppm (58.3%). Lower concentrations of
GA3 were less effective. For H. petiolaris, the most effective concentrations of GAS were 1000 ppm
(56.1%), 500 ppm (65.0%), and 250 ppm (62.2%) and, again, lower concentrations of GAS were less
effective. Transplanting the germinated seeds of H. annuus to soil in pots, resulted in nearly 100%
success, indicating no apparent long-term effects from the GAS treatment. Published on-line
www.phytologia.org P/nto/og/fl 99(1): 32-35 (Jan 19, 2017). ISSN 0S0S194S0.
KEY WORDS: Helianthus annuus, H. petiolaris, seed germination, dormancy, gibberellic acid (GAS).
Native, wild sunflowers (Helianthus spp.) are known to be difficult to germinate (Seiler, 199S).
Recently, Adams and TeBeest (2016) reported on various stratification treatment effects on germination
of Helianthus petiolaris. We found a moderate concentration of GAS (500 ppm) with one week
stratification at 4°C was very effective in increasing the germination rate of recalcitrant native sunflower
seeds (80% vs. S0% control). Stratification (1 wk at 4°C) increased germination, regardless of the seed
treatment. Ethrel (25 ppm) treatment was effective, but not as much as GAS (500 ppm). Soaking
sunflower seed in water for 12 or 16 hr resulted in no seed germination.
The literature on pre-treatment methods for sunflower seed germination has been recently
reviewed (Adams and TeBeest, 2016).
The purpose of the present paper is extend the tests on pre-treatment using GAS at different
concentrations to determine the concentration of GAS that produces highest seed germination in H
annuus and H. petiolaris, native, wild collected seed.
MATERIALS AND METHODS
Seeds of H petiolaris, PI451978-NC7, Ellsworth, KS were obtained from GRIN (Germplasm Resources
Information Network), USDA.
Seeds of H. annuus: were collected 16 July 2016, from a natural population, 1 mi. south of Gruver, TX
(Adams 14952).
All seeds were surface sterilized by:
1. Washing with soap/tap water;
2. Dipping in 70% ethanol, SO sec;
S. Sterilizing by soaking in 20% Chlorox (8.25% sodium hypochlorite) for SO min.;
4. Thoroughly rinsing in sterilized ddwater (Protocol from Singhung Park, Kansas State University).
Phytologia (Jan 19, 2017) 99(1)
33
Germination tests: Effects of various concentrations of gibberellic acid (GA3, PlantHarmones.net, 90%)
stored at 4°C, 1 week (7 days) in GA3 solutions.
1000 ppm GA3 stock solution: dissolved 1.0 g GA3 in 5 ml ethanol, added to 995 ml DI water to produce
1000 ppm stock. Diluted with DI water to make: 500 ppm, 250 ppm, 125 ppm, and 62.5 ppm stocks.
Control: soaked in DI water, 4°C, 1 week.
20 seeds were used in each of 3 replicates (60 seeds total). The seeds were soaked in DI, or various GA3
solutions in beakers, 4°C, 1 week. In addition, for both H. annuus and H. petiolaris, 60 seeds were placed
in sterilized filter paper, pre-wetted with 500 ppm GA3, then placed in sealed plastic bags at 4°C, 1 week.
Seeds were germinated at RT (21°C), in normal lab fluorescent lighting. Seeds were examined for fungal
contamination daily and contaminated seeds removed. After 14 days, the seeds with emergent roots were
scored as germinated.
RESULTS
Table 1 shows that for H. annuus, the most effective concentrations of GA3 were 1000 ppm
(61.7%) and 500 ppm (58.3%). Lower concentrations of GA3 were less effective. This is shown in
figure 1, where 1000 and 500 ppm were much more effective than lower concentration of GA3.
c
g
‘t— ■
03
c
’£
CD
control 1000 ppm 500 ppm 250 ppm 125 ppm
Pre-Treatments
Figure 1. Germination of H. annuus at various concentrations of GA3, soaked 1 week, 4°C. Control:
soaked in DI water, 1 week, 4°C.
For H. petiolaris, the most effective concentrations of GA3 were 1000 ppm (56.1%), 500 ppm
(65.0%), and 250 ppm (62.2%).
In contrast to the results for H. annuus, lower concentration of GA3 were somewhat effective in
seed germination for H. petiolaris (Fig. 2), with considerable enhanced germination at 250 and 125 ppm
GA3.
Comparing soaking seeds in a beaker of 500 ppm GA3 vs. storage in filter paper saturated with
500 ppm GA3 resulted 58.3% vs 51.7% (H. annuus. Table 1) and 65.0% vs. 44.8% (H. petiolaris. Table
2). This seems to indicate that there is a small advantage in soaking the seeds in a beaker.
34
Phytologia (Jan 19, 2017) 99(1)
H. petiolaris
70
60
Cl
o
50
15
c
40
E
t
Q
05
30
20
10
0
control 1000 ppm
500 ppm 250 ppm
Pre-Treatments
125 ppm 62.5 ppm
Figure 1. Germination of H. petiolaris at various concentrations of GAS, soaked 1 week, 4°C. Control
soaked in DI water, 1 week, 4°C.
In summary, this study found an effective pre-treatment to enhance seed germination of H.
annuus and H. petiolaris is soaking in GAS for 1 week, 4°C. It should be noted that transplanting
germinated seeds of H. annuus to soil in pots, resulted in nearly 100% success, indicating no apparent
long-term effects from the GAS treatment.
ACKNOWLEDGEMENTS
This research funded by Baylor University. Thanks to Laura Marek and Lisa Pfiffner, GRIN,
USD A, for helpful discussions
LITERATURE CITED
Adams, R. P. and A. K. TeBeest. 2016. The effects of gibberellic acid (GAS), Ethrel, seed soaking and
pre-treatment storage temperatures on seed germination of Helianthus annuus and H. petiolaris.
Phytologia 98: 213-218.
Kumari, C. A. and B. G. Singh. 2000. Ethephon adequacy in release of innate dormancy in sunflower.
Indian J. Plant Physiol. 5: 277-280.
Maiti, R. K., P. Vidyasagar, S. C. Shahapur and G. J. Seiler. 2006. Studies on genotype variability and
seed dormancy in sunflower genotypes (Helianthus annuus L.). Indian J. Crop Sci. 1: 84-87.
Seiler, G. J. 1993. Wild sunflower species germination. Helia 16: 15-20.
Phytologia (Jan 19, 2017) 99(1)
35
Table 1. Germination tests of H. annuus, native, Gruver, TX. References: Kumari and Singh (2000)
Maiti et al. 2006.
Pre-Treatment, all soaked 1 wk, 4°C
germination rates
1. control: seeds soaked in DI water
9/60 = 16.7%
2. 1000 ppm GA3
37/60-61.7%
3. 500 ppm GA3
35/60 - 58.3%
3a. 500 ppm GA3 on filter paper
31/60-51.7%
4. 250ppm GA3
15/60 - 25.0%
5. 125ppm GA3
14/60 - 23.2%
Table 1. Germination tests of H. petiolaris, native, Ellsworth, KS. References: Kumari and Singh (2000)
Maiti et al. 2006.
Pre-Treatment, aU soaked 1 wk, 4°C
germination rates
1. control: seeds soaked in DI water
6/59 -10.2%
2. 1000 ppm GA3
32/57-56.1%
3. 500 ppm GA3
39/60 - 65.0%
3a. 500 ppm GA3 on filter paper
26/58 - 44.8%
4. 250 ppm GA3
33/53 - 62.8%
5. 125 ppm GA3
28/57-49.1%
6. 62.5 ppm GA3
21/58 - 36.2%
36
Phytologia (Jan 19, 2017) 99(1)
Legitimacy of the name Croton bigbendensis (Euphorbiaceae)
Billie L. Turner
Plant Resources Center, The University of Texas, Austin, TX 78712
billie.turner@austin.utexas.edu
ABSTRACT
The legitimacy of the name Croton higbendensis is discussed and the circumstances concerning
the issuance of a Holotype based on pistillate and staminate plants explained. Published on-line
www.phytologia.org Phytologia 99(1): 36-37 (Jan 19, 2017). ISSN 030319430.
KEY WORDS: Croton higbendensis, nomenclature, holotype.
Turner (2004) published the name Croton bigbendensis B.l. Turner, this largely confined to the
southern Big Bend region of western Texas. The taxon was typified by a single collection (composed of
several plants) at the same place at the same time. Because the population was composed of both
pistillate and staminate plants, I provided the number Turner 22-204A for the pistillate plants and Turner
22-204b for the staminate plants. The plants concerned clearly belonged to the same collection, all
bearing the same number, although I did designate a pistillate plant from the population as the Holotype,
however, my intent was to treat Turner 22-204 (both A and b) as holotype material, this clearly stated and
so pictured in my figures 1 and 2. But some purists (cf. discussion provided by Wiersema 2016) view
such typification as contrary to the Code, contending that only a single plant number should have been
applied to the Holotype, thus invalidating the name, although my application of such was quite clear, this
discussed further in more detail by my archrival, Henrickson (2010), who would recognize my novelty as
but a variety, at best, this after a lengthy digression into my systematic mores.
Strangely, W. van Ee and Berry (2016), did not account for the name C. bigbendensis in their
treatment of Croton for the Flora of North America, nor did they mention the work of Henrickson. I
would like to place on record here that I believe the name C. bigbendensis B.L. Turner is properly
typified and deserves recognition, as justified in the above. As to the taxonomic criticism of the taxon
posited by Henrickson, I leave such evaluation to future workers having not the bias Henrickson and I
both possess.
An up to date distributional map of C. bigbendensis is provided in the present account (Fig. 1),
this part of my Atlas of the Vascular Plants of Texas (Turner 2017, in prep.).
LITERATURE CITED
Ee, W. van and P.E Beny. 2016. Croton, in N. Amer. FI. 12: 206-224.
Henrickson, J. 2010. Croton bigbendensis Turner (Euphorbiaceae) Revisited. J. Bot. Res. Inst. Texas 4:
295-301.
Turner, B.L. 2004. Croton bigbendensis (Euphorbiaceae), anew species from Trans-Pecos, Texas. SIDA
21:79-85.
Turner, B.L. 2017. Atlas of the Vascular Plants of Texas [2"^^ edition, in prep].
Wiersema, J.H. 2016. Proposal to provide a more direct definition of the term “gathering". Taxon 65:
1186.
Phytologia (Jan 19, 2017) 99(1)
Figure 1. Distribution of Croton bigbendensis in Texas.
38
Phytologia (Jan 19, 2017) 99(1)
Multiple evidences of past evolution are hidden in nrDNA of Juniperus arizonica and
J. coahuilensis populations in the trans-Pecos, Texas region
Robert P. Adams
Biology Department, Baylor University, Box 97388, Waco, TX 76798, USA, Robert_Adams@baylor.edu
ABSTRACT
Geographical analysis of variation in nrDNA polymorphisms of J. arizonica and J. coahuilensis
in the trans-Pecos, TX region showed multiple patterns of hybridization, both modem and relictual
(Pleistocene) introgression, incomplete lineage sorting and relictual hybridization. The concept that
nrDNA from a single plant could harbor multiple evidences of past evolution appears to be novel. Total
nrDNA polymorphisms were maximal in the Ft. Davis, Alpine, Marfa trans-Pecos area and on the
granitic rocks at Hueco Tanks State Park, TX. Published on-line www.phytologia.org Phytologia 99(1):
38-47 (Jan 19, 2017). ISSN 030319430.
KEY WORDS: Juniperus arizonica, J. coahuilensis, Cupressaceae, hybridization, introgression,
incomplete lineage sorting, nrDNA polymorphisms, petN-psbM DNA.
Recently, Adams (2016) found (by petN-psbM sequencing) that Juniperus arizonica, previously
known only from Arizona and New Mexico, occurs in trans-Pecos Texas in the Fra nklin Mtns., Hueco
Mtns., Hueco Tanks State Park, Quitman Mtns., Eagle Mtns. and Sierra Vieja Mtns., primarily on igneous
material (Figs. 1, 2). These trans-Pecos juniper populations have previously been identified as J.
coahuilensis . These taxa have very distinct differences in their DNA and are in separate clades (Adams,
2014, Adams and Schwarzbach, 201 1, 2013). The cp region petN-psbM is especially efficient in
separating these taxa, as 5 SNPs occur in the 794 bp region.
Figure 2. Distribution of J. coahuilensis
Detailed mapping of plants by their cp DNA (petN-psbM) showed that all the plants (or
specimens) in New Mexico and northern Mexico, as well as plants examined from the Franklin Mts.,
Hueco Tanks SP, Quitman Mts., Eagle Mts., and one plant from Sierra Vieja Mtns. contained the J.
arizonica cp DNA (Fig. 3). Junipers from Ft. Davis, Alpine, Marfa and Big Bend (1) all had the J.
coahuilensis cp DNA (Fig. 3). The occurrence and extent of hybridization and introgression in that
Phytologia (Jan 19, 2017) 99(1)
39
region is not known, except for a study of hybridization between J. coahuilensis and J. pinchotii in the
Chisos Mtns. (Adams and Kistler, 1991).
Recently, Adams et al. (2016) have reported that in the sister genus, Hesperocyparis, artificial
hybrids between Hesperocyparis (= Cupressus in part) arizonica and H. macrocarpa, nrDNA was
inherited as heterozygous for diagnostically different sites. They concluded that, at least in
Hesperocyparis (and likely in the Cupressaceae, including Juniperus), analysis of heterozygous nrDNA
(ITS) could be used for the detection and analysis of hybridization. Because F 2 progeny and backcrosses
were not analyzed, they could not comment on the amount and/ or speed of lineage sorting in
Hesperocyparis.
The purpose of this paper is to report on the composition of nrDNA in populations in the trans-
Pecos, TX region and the investigation of hybridization, introgression and incomplete lineage sorting.
Figure 3. Distribution of J. arizonica and J. coahuilensis based on petN-psbM cp data.
MATERIALS AND METHODS
Plant material and populations studied:
Sedona, AZ
J. arizonica by petN DNA, common in grassland, tree 6 m tall, female, with J. osteosperma on alluvial
soil. On AZ highway 179, between Sedona and 117. 34° 42.43rN, 111° 46.369’ W, 1150 m, 13 March,
2005, Yavapai Co., AZ, Robert P. Adams 10634-10636,
Cottonwood, AZ
J. arizonica by petN DNA, abundant, on alluvial fan, 3 mi. SW of Cottonwood, AZ, on D. Thornburg's
property, 34° 41' 17.4" N, 112° 03' 05.46" W, 4060 ft., 13 Jan., 2010, Yavapai, Co., AZ, Coll. David
Thornburg ns. Lab Ace. Robert P. Adams 14908-14913,
40
Phytologia (Jan 19, 2017) 99(1)
Southern New Mexico:
J. arizonica by petN DNA, Hidalgo Co., NM, Animas Mtns, 31.61176° N, 108.7791° W, 5750', Seinet
Cat# 57778, Wagner 1283. 22 Jul 1975, Lab Acc. Robert P. Adams 14697,
7. arizonica by petN DNA, Luna Co., NM, Tres Hemianos Mtns, 31.9010° N, 107.7794° W, 4250', Seinet
Cat # 85666, 7 L Carter 1246, 14 Aug 1993, Lab Acc. Robert P. Adams 14698,
7. arizonica by petN DNA, Hidalgo Co., NM. Animas Mtns, 31.5938° N, 108.7684° W, 6000', Seinet Cat
# 57776, Wagner 1005, 17 Jun 1975, Lab Acc. Robert P. Adams 14701,
7. arizonica by petN DNA, no cones, Hidalgo Co., NM, Animas Peak, Animas Mtns., 31.5813° N,
108.7843° W, 8452' (Google Earth), Seinet Cat# 25131, WC Martin 4678, 29 Oct 1960, Lab Acc.
Robert P. Adams 14705,
7. arizonica by petN DNA, Hidalgo Co., NM, Big Hatchet Mtns.,with Quercus, Parthenium, Ocotillo,
Mesquite, Agave 31.6249° N, 108.36425° W, 5350', Ken Heil 9254, 28 May 2010, Lab Acc. Robert P.
Adams 14716,
7. arizonica by petN DNA, Grant Co., NM, ca 1 .5 mi. s of NM hwy 9, near 'Old Hachiti' townsite.
Chihuahuan desert scrub - creosote, Lycium koberlina and Dalea formosa. 31.9139° N, 108.41472° W,
4745', Ken Heil 32357, 29 Apr 2010, Lab Acc. Robert P. Adams 14717,
Rock Hound State Park, NM (type locality, 7. arizonica)
7. arizonica by petN DNA, multi-stemmed shrubs to 4m, in Bouteloua grassland. Pollen shed in Mar-
April?, Fruit rose color. Rock Hound State Park. 17km S, and 8 south of Deming, NM, 32° 1 l.lbl’N,
107° 36.651’ W, 1420 m, 6 Feb., 1996, Luna Co., NM„ Robert P. Adams 7635-7637
7. arizonica by petN DNA, common in Bouteloua grassland, shrub-trees to 3-5 m.. Rock Hound State
Park., 32° 1 L161’N, 107° 36.651’ W, 1420 m, 12 Mar, 2005, Luna Co., NM, Robert P, Adams 10630,
Quitman Mtns.
7. arizonica by petN, Hudspeth Co., TX, common on degraded granite, north face of Quitman Mtns., with
desert-scrub. On .south .side of 110, ~6.3 mi. w of Sien’a Blanca, TX, 31°12' 25" N; 105° 27' 51" W,
4629’, Robert P. Adams 14798-14806, 12 March 2016,
Hueco Tanks St. Park, TX
7. arizonica by petN El Paso Co., TX, uncommon, 50- 100 trees seen, on granite, Hueco Tanks St. Park,
31° 54’ 49.7" N; 106° 02' 6.8" W, 4560', Robert P. Adams 14827-14835, 18 March 2016. Robert P.
Adams 14718.
11.2 s of Alpine, TX on Tex 118
7. coahuilensis, by petN, Brewster Co, TX, abundant in grassland, 11.2 s of Alpine, TX on Tex 118. 30°
14' 08" N; 103° 34' 00" W, 5222', Robert P. Adams 14807-14811, 15 March 2016,
11.0 mi w of Alpine, TX on US 90
7. coahuilensis, by petN, Brewster Co, TX, 1 1,0 mi w of Alpine on US 90, abundant in grassland, in
Paisano Mtns., 30° 17' 42" N; 103° 48' 02" W, 4967’, Roben P. Adams 14812-14816,^5 March 2016,
4.2 mi se of Ft, Davis, TX on Tex 118, CDRI
7. coahuilensis, Jeff Davis Co., TX, common locally, in grassland. 4.2 mi se of Ft. Davis, on Tex 118, e
1 .0 mi into Chi, Desert Res. Inst., 39° 09' 27.54" N; 86° 18' 23.31" W, 5050', Robert P. Adams 14817-
14821, 16 March 2016,
19.4 mi. s of Marfa, TX on US 67
7. coahuilensis, bv petN, Presidio Co., TX, common in grassland, 19.4 mi. s of Marfa, on US 67, 30° 04'
07" N; 104° 10'" 19" W, 5137', Robert P. Adams 14822-14826, 16 March 2016,
La Zarca, Mexico
7. coahuilensis, large population with thousands of trees. 85 km N. of La Zarca on Mex. 45, 1740m, 10
Dec, 1991, Durango, Mexico, Robert P. Adams 6829-6831,
Voucher specimens for new collections are deposited in the Herbarium, Baylor University (BAYLU).
One gram (fresh weight) of the foliage was placed in 20 g of activated silica gel and transported
to the lab, thence stored at -20° C until the DNA was extracted. DNA was extracted from juniper leaves
by use of a Qiagen mini-plant kit (Qiagen, Valencia, CA) as per manufacturer’s instructions.
Phytologia (Jan 19, 2017) 99(1)
41
Amplifications were performed in 30 pi reactions using 6 ng of genomic DNA, 1.5 units Epi-
centre Fail-Safe Taq polymerase, 15 pi 2x buffer E (petN-psbM), D (maldehy) or K (nrDNA) (final
concentration: 50 mM KCl, 50 mM Tris-HCl (pH 8.3), 200 pM each dNTP, plus Epi-Centre proprietary
enhancers with 1.5 - 3.5 mM MgCl 2 according to the buffer used) 1.8 pM each primer. See Adams, Bartel
and Price (2009) for the petN-psbM primers utilized.
The PCR reaction was subjected to purification by agarose gel electrophoresis. In each case, the
band was excised and purified using a Qiagen QIAquick gel extraction kit (Qiagen, Valencia, CA). The
gel purified DNA band with the appropriate sequencing primer was sent to McLab Inc. (San Francisco)
for sequencing. Sequences for both strands were edited and a consensus sequence was produced using
Chromas, version 2.31 (Technelysium Pty Ltd.).
RESULTS AND DISCUSSION
Sequencing petN-psbM yielded 794 bp with 5 SNPs separating J. arizonica and J. coahuilensis.
In addition, nrDNA was sequenced yielding 1270 bp with only 1 SNP (at site 533) separating J. arizonica
and J. coahuilensis. Using these data, samples were classified accordingly (Table 1). Based on
heterozygous peaks at site 533, 11 samples were classified as hybrids (AxC, Table 1). According to
nrDNA, hybrids occur mostly in the Anim as Mts., NM, Hueco Tanks SP, TX and Quitman Mtns., TX
(Fig. 4.). Note one hybrid in the Marfa, TX population. The nrDNA data, indicates that populations of J.
coahuilensis in the Alpine - Ft. Davis - Marfa area are nearly pure. It should be noted that the soils of
Hueco Tanks and Quitman Mtns. are granitie, whereas the Alpine - Ft. Davis - Marfa area soils are
volcanic.
Fig. 4. Distribution of J. arizonica x J. coahuilensis hybrids based on nrDNA.
42
Phytologia (Jan 19, 2017) 99(1)
In order to visualize the correlation of nrDNA and cp (petN) classifications, each plant was
scored for species or hybrid for nrDNA and cpDNA. Mapping this classification shows a relatively sharp
demarcation between J. arizonica and J. coahuilensis (Fig. 5). The zone of hybridization is in Hueco
Tanks State Park, Quitman Mtns., and Anima Mtns. and this appears to be a region of introgression
northward from J. coahuilensis (Fig. 5).
The Hueco Tanks State Park, Quitman Mtns., and Anima Mtns. populations are on granitic soil
and the J. coahuilensis populations in the Ft. Davis, Alpine, Marfa region are on volcanic soil. Soil
differences may be the factor that determines the northern range of J. coahuilensis and could present a
barrier for additional introgression northward into J. arizonica.
Figure 5. Mapping plants showing their classification as J. arizonica, J. coahuilensis, or hybrids for both
nrDNA and cpDNA.
Mapping the number of nrDNA polymorphic sites per plant shows very low polymorphic sites in
the normal range of J. arizonica (NM and AZ, Fig. 6). However, where J. arizonica and J. coahuilensis
hybridize and thence southward, there are several populations with plants having 1 to 6 polymorphic sites
(excluding site 533). Hueco Tanks is very variable: 3 plants with 0 polymorphisms; 3 with 1; 2 with 5;
and 1 with 6 polymorphisms (Fig. 6). The Davis Mtns - Alpine area is also a region with lots of
polymorphisms (Fig. 6). In contrast to the more mountainous sites, the Marfa population (19.3 mi sw of
Marfa, in a Bouteloua grassland) had low polymorphisms in its nrDNA.
Phytologia (Jan 19, 2017) 99(1)
43
The trans-Pecos region likely experienced a mixing of southern Rockies flora to move southward
and the flora of the Sierra Madre Oriental flora to move northward during cooling and heating eras in the
Pleistocene. This provided opportunities for many Juniperus species, now spatially separated, to
hybridize in the past.
Fig. 6. Distribution of the number of nrDNA polymorphisms/ plant.
A closer examination of individual plant nrDNA site polymorphisms revealed that nrDNA
harbors several evolutionary patterns that vary by region. For site 543, 1 1 plants contained (C/G) and
these range from the Quitman Mtns., northwest to Cottonwood and Sedona, AZ (Fig. 7). Only one plant
was G/G, and that was in the Cottonwood, AZ population. This is near the northwestern limit of J.
arizonica. Site 543 might be an indicator of introgression from J. coahuilensis into J. arizonica.
In addition, another polymorphism occurs (C/T, Fig. 7), but only in the La Zarca, MX population.
Additional research is needed to determine if the T comes from hybridization with another Mexican
juniper, from incomplete lineage sorting or just a local mutation.
44
Phytologia (Jan 19, 2017) 99(1)
The distribution of
variation in site 173 (A/G) is
centered between J. arizonica
and J. coahuilensis in the
Animas Mtns., NM, Hueco
Tanks SP, and Quitman Mtns.
(Fig. 8.) This may to be either
relictual hybridization, or
incomplete lineage sorting.
nrDNA site 304
contains two geographical
patterns. One (A/T, Fig. 9) is
similar to that for site 173 (Fig.
8) in the Animas Mtns., NM,
Hueco Tanks SP, and Quitman
Mtns. The second pattern (C/T,
Fig. 9) is found in only the south
Alpine, TX population. The C/T
site might be due to
introgression from the east or
south from Mexico, perhaps
from mixing of taxa during the
Pleistocene. Or it may be just a
local mutation in that population.
Figure. 7. Distribution of polymorphisms at nrDNA site 543.
Figure 8. Dist. of nrDNA site 173 polymorphism. Figure 9. Dist. of nrDNA site 302 variation.
Phytologia (Jan 19, 2017) 99(1)
45
Variation in site 318 (C/T, Fig. 10) spans
the J. arizonica - J. coahuilensis range junetion
and seems likely to be from relictual
hybridization. There appears no source of the C
allele in any population of J. arizonica or J.
coahuilensis examined. Alternatively, it could be
incomplete lineage sorting.
Finally, two sites show very similar
patterns: both sets of polymorphisms are confined
to the Ft. Davis - Alpine - Marfa area and both
sites have plants with mixed bases as well as
plants with homozygous bases. Site 302 (A/G)
was found in all four populations, plants
homozygous for A are in all 4 populations, but
only one plant homozygous for G was found (in
the Ft. Davis population. Fig. 11).
polymorphisms.
A similar pattern was found for site 303 (C/T). C/T was present in all four populations, plants
homozygous for C were in all 4 populations, but only 2 plants homozygous for T occurred in the Ft.
Davis and Marfa populations (Fig. 12). These two sites are difficult to explain. It almost appears that an
unknown (to the author) species is present that has (G,C) at 302, 303 and is hybridizing with J.
coahuilensis (A, C) at 302,303. Other juniper species in the area are J. pinchotii (Kent, and Fort
Stockton), J. monosperma (near Kent), and J. deppeana (higher elevations in the area). Of course, it
might be Pleistocene relictual hybridization with a species (or its ancestor) now growing in Mexico.
Figure 11. Geographical variation in variation at Figure 12. Variation in nrDNA site 303.
nrDNA site 302.
46
Phytologia (Jan 19, 2017) 99(1)
SUMMARY
Geographical analysis of variation in nrDNA polymorphisms of J. arizonica and J. coahuilensis
in the trans-Pecos, TX region showed multiple patterns of hybridization, both modem and relictual
(Pleistocene) introgression, incomplete lineage sorting and relictual hybridization.
The concept that nrDNA from a single plant could harbor multiple evidences of past evolution
appears to be novel. The pre-occupation of evolutionary systematists with phylogeny has resulted a lack
of critical variation in nrDNA. Heretofore, the standard procedure is to sequence nrDNA (as the sole
proxy of the nuclear DNA), then add in a few cpDNA sequences, then ran the data in a phylogenetics
software and publish 'the Phylogeny', and then move on to another genus. That may satisfy a need for a
broad evolutionary framework of a group (genus). But, as shown in this report, there may be
considerable evidence of past evolutionary events in nrDNA that would be completely ignored (and
missed) by only running a phylogenetic analysis.
Total nrDNA polymorphisms were maximal in the Ft. Davis, Alpine, Marfa trans-Pecos area and
on the granitic rocks at Hueco Tanks State Park, TX. Additional research using Single Copy Nuclear
Genes (SCNG) is needed to further address the variation found in this region.
ACKNOWLEDGEMENTS
Thanks to George M. Ferguson (UA), Ken Heil (SJNM), Tim Lowrey (UNM), Mike Powell
(SRSC) and Richard Worthington (UTEP) for letting me sample (or sending small fragments) herbarium
specimens. This research was supported in part with funds from Baylor University. Thanks to Amy
TeBeest for lab assistance and Andrea Schwarzbach for helpful suggestions on the manuscript.
LITERATURE CITED
Adams, R. P. 2014. The junipers of the world: The genus Juniperus. 4th ed. Trafford PubL, Victoria, BC.
Adams, R. P. 2016. Juniperus arizonica (R. P. Adams) R. P. Adams, new to Texas. Phytologia 98: 179-
185
Adams, R. P. J. A. Bartel and R. A. Price. 2009. A new genus, Hesperocyparis, for the cypresses of the
new world. Phytologia 91: 160-185.
Adams, R. P. and J. R. Kistler. 1991. Hybridization between Juniperus etythrocarpa Cory and Juniperus
pinchotii Sudworth in the Chisos Mountains, Texas. Southwest. Natl. 36: 295-301.
Adams, R. P,, M. Miller and C. Low. 2016. Inheritance of nrDNA in aitificial hybrids of Hesperocyparis
arizonica x H. macrocarpa. Phytologia 98: 277-283.
Adams, R. P. and A. E. Schwarzbach. 201 1 . DNA barcoding a juniper: the case of the south Texas
Duval county juniper and sen'ate junipers of North America. Phytologia 93(1): 146-154.
Adams, R. P. and A. E. Schwarzbach. 2013. Taxonomy of the seiTate \eSLf Juniperus of North America:
Phylogenetic analyses using nrDNA and four cpDNA regions. Phytologia 95: 172-178.
Phytologia (Jan 19, 2017) 99(1)
47
Table 1. Classification of samples based on petN and nrDNA. Bold are putative hybrids between J. arizonica and J. coahuilensis
by ITS site 533, A in arizonica, T in coahuilensis, (A/T in 533) were scored as hybrids (AxC). The more common polymorphic
605 1 708 I# poly
sites are shown. A few rarer,
polymorphic site
Sample
ITS
az 1 0634Sedona 1 8 1 M
ariz
ariz
azl0635Sedoiia 68 IM
ariz
ariz
azI0636Sedona
aiiz
aiiz
az 14908Cotlonwood
ariz
ariz
az 1 4909Cottonvvood
ariz
ariz
az 1 49 1 OCottoii wood
ariz
ariz
az 1 49 1 2C(>lionwood
ariz
ai'iz
az 1 49 1 3Cotton wood 1 2 1 Y
ariz
ariz
azl4717GrantCoNM 18IM
ariz
ariz
az7635RockIToundSP
ariz
ariz
az 7 636RockH o undSP
ariz
ariz
az7637RockHoiindSP
ariz
ariz
az 10630RockHoundSP
ariz
ariz
az 14698LunaCoNM
ariz
ariz
azl4697HidaleoCoNM
ariz
ariz
azI470lHidalaoCoNM
ariz
ariz
azl47U5HdalgoCoNM
ariz
AxC
azl4716HidalgoCoNM
ariz
AxC
coal4827HuecoTanks
ariz
coah
coa 1482X11 iii'coTanks
ariz
AxC
coa 1482911 uecoT anks
ariz
AxC
coa 14830HuecoTanks
ariz
AxC
coal4831HuecoTanks
ariz
AxC
coa 1 48 32HuecoTanks
ariz
coah
coa 14833H uecoTanks
ariz
AxC
coa 1 4834HuccoTanks
ariz
ariz
coa 1 4835HuecoTanks
ariz
ariz
coal4798 Quitman Mtns-
ariz
AxC
coa 14799 Quieman Mtns.
ariz
ariz
coa 1 48CiCi Q ui tman M tn s .
ariz
aiiz
coal4S01 Quitman Mtns.
ariz
AxC
coa 14802 Quitman Mtns.
ariz
ariz
coal4803QtiitmanMtns.804R
ariz
AxC
coal4804Quuman Mtns. 804R
ariz
coah
coa 14805 Quitman Mtns.
ariz
ariz
coa 1 4 8 06Q ui tman Muis .
ariz
ariz
coa 14807sorAlpme
coah
coah
coa 1 4808sOtAlpine
coah
coah
coa 1 48 1 Ospf Alpine
coah
coah
coa 14811 so f Alpine
coah
coah
coal 48 1 2wofAlpine
coah
coah
coal48]3wafAlpine 313R
coah
coah
coa 1481 4wor.\lpine
coah
coah
coal48 1 SwofAlpine
coah
coah
coa 1 48 1 6vvotAlpine
coah
coah
coal4S17FiDavis lOOOY
coah
coah
coal4818FtDavis
coah
coah
coal48l9FtDavis
coah
coah
coal4820FtDavis 689K
coah
coah
coal 4821 FtDav is llOOY
coah
coah
coa 1 4822sot'Marfa
coah
coah
coal4823sotMaifal lOOY
coah
coah
coa 1 4824solMarra
coah
coah
coa 14825s«f \ larfa
coah
AxC
coal4826sol'MarfaY1100
coah
coah
coa6829L;iZarca
coah
coah
coa6830LaZarca
coah
coah
coa683 1 LaZatca
coah
coah
coa 1 024 1 knj45nDgo
coah
coah
coal 0242k m45nDgo,503Y
coah
coah
number of polymorphic
48
Phytologia (Jan 19, 2017) 99(1)
Comparison of leaf essential oils of fastigiate (strict) and horizontal forms of
Cupressus sempervirens from Cyprus, Montenegro, Turkey, and United States.
Robert P. Adams
Biology Department, Baylor University, Box 97388, Waeo, TX
76798, USA,
Robert_Adams@baylor.edu
Tugrul Mataraci
Eskidji Mtiz. A§., Sanayi Cad. Vadi Sokak No: 2, Tarabya, Istanbul, Turkey
Salih Gticel
Environmental Researeh Institute, Near East University, North Nieosia, Cyprus
and
Jim A. Bartel
San Diego Botanie Garden, P. O. Box 230005, Eneinitas, CA 92023
ABSTRACT
The volatile leaf oils of the horizontal form of C. sempervirens from natural populations in
Cypms and Turkey were very unifomi and dominated by a-pinene (36.2, 26.0%), myrcene (2.4, 2.4%), 6-
3-carene (18.3, 16.0%), terpinolene (3.2, 3.8%), a-terpinyl acetate (4.7, 3.5%), cedrol (4.4, 3.3%), manoyl
oxide (0.7, 3.8%), iso-pimara-7, 15-diene (0.4, 2.6%), isoabienol (2.4, 4.0%), and trans-totarol (1.5,
5.7%). Overall, the major terpenes compositions were very uniform for the sampled natural populations
(Cypms, Turkey) and fastigiate (strict) fonns from California and Istanbul. But they were very variable
for the oils from other fastigiate forms (Turkey and Montenegro). The fastigiate forms of Cupressus
sempervirens from California and Istanbul (14674) have oils that are similar to natural populations.
Variation in the composition of oils from cultivated fastigiate fonns in Turkey and Montenegro suggests
that these cultivars arose from multiple selections of fastigiate (strict) trees, rather than cloning and
widespread cultivation. The volatile leaf oil composition does not support the recognition of the tvso
growth forms of C. sempet^irens as distinct taxa. Published on-line mvw.phytologia.org Phytologia
99(1): 48-53 (Jan 19, 2017). ISSN 030319430.
KEY WORDS; Cupressus sempervirens, C. horizontalis, C. fastigiata, terpenoids, geographic variation,
taxonomy.
Cupressus sempervirens E. ranges naturally from the eastern Mediterranean, Crete, Cyprus,
eastern Aegean Islands, Iran, Israel, Jordan, Lebanon, Syria, Turkey, and possibly Libya (S^kiewicz et al.
2016). The species has been widely cultivated within and outside its range throughout the warm
temperate world (More and White 2002). Farjon (2005, 2010) noted that C. sempervirens has
traditionally been separated into two "‘elements”; pyramidal trees with horizontal branches (horizontal
fonu) (= C. horizontalis Milk) and fastigiate trees with strict branching (fastigiate form) (= C. fastigiata
DC.). The fastigiate trees are often called Italian, cemetery, graveyard, and Tuscan cypress in the Old
World, while m tire New World, the widely cultivated fastigiate cultivars are called Italian and cemetery
cypress. Faijon (2005) concluded that the fastigiate (strict) form of C. sempei'virens, widely cultivated all
over the Mediterranean and beyond, was selected many centuries ago from natural populations, which
likely were largely horizontal.
Phytologia (Jan 19, 2017) 99(1)
49
The volatile leaf essential oils of Cupressus sempervirens (both horizontal and fastigiate forms)
have been analyzed based mostly on locally cultivated fastigiate trees. The report by Ulukanli et al.
(2014) is typical reporting the major components being: a-pinene (35.6%), trans-pinocarveol (5.22%), a-
phellandrene-8-ol (4.56%), f3-pinene (3.1%), limonene (2.8%), bonieol 2.3%) and camphene (2.2%).
Chanegrilia, et al. ( 1977) reported on the leaf oils of C sempervirens from Algeria (cv. strictaJ) as having
a-pinene (44.9%), 6-3-carene (TO. 6%), limonene (4.5%), terpinolene (2.7%), terpin-4-ol (T.9%), a-
terpinyl acetate ( 12.0%) and rnanoyl acetate (1,5%). Florcani et al. (1981) reported the essential oil of cv.
stricta (Argentina) contained a-pinene (50.1%), camphene (1.4%), p-pinene (4.1%), 6-3-carene (30.5%),
limonene (3.5%), terpinolene (1.3%) and a-terpineol (1.6%). Other reports are by Adams et al. (1997),
Amri et al. (2013), Pauly et al. (1983), Floreani et al. (1982), and Gamero et al. (1978)
This paper compares the volatile leaf oil of the horizontal fonn of C. sempennrens from natural
populations in Cyprus and Turkey to that of cultivated fastigiate trees from Montenegro, Turkey, and
California, USA.
MATERIALS AND METHODS
Plant materials:
Cupressus sempervirens L. (horizontal form):
Cypru.s: 35° 16' 34.58" N, 33° 23' 14.12" E, 361 m, 3 June 2015, Salih Gucel ns, Lab Acc. Robert P.
Adams 4560-14564,
Turkey: pyramidal trees, branches horizontal,
Vicinity of Beskonak village, Serik, Antalya, 37° 17' N, 31° 18' E, elev. 180 m, 23 May 2015, Coll.
Tugrul Malaraci, 2015-14, Lab Acc: Robert P. Adams 14565,
In Koprulu Kanyon National Park, on the road of Ancient city of Selge, Beskonak village, Serik,
Antalya, 37° 21' N, 31° 53' E, elev. 708 m, 23 May 2015, Coll. Tugrul Mataraci, 2015-15, Lab. Acc:
Robert F. Adams 14566.
In Kdpriilu Kanyon National Park, on the road of Ancient city of Selge, Beskonak village, Serik,
Antalya, 37° 22' N, 3 1 ° 1 3' E, elev. 817m, 23 May 20 1 5, Coll. Tugrul Mataraci, 2015-16, Lab. Acc:
Robert P. Adams 14567.
In Kdprulii Kanyon National Park, on the road of Ancient city of Selge, Beskonak village, Serik,
Antalya, 37° 21' N, 3 1° 14' E, elev. 764 m, 23 May 2015, Coll. Tugrul Mataraci, 2015-17, Lab. Acc:
Robert P. Adams 14568
Cupressus sempervirens (fastigiate form):
Montenegro:
fastigiate (strict), columnar tree in maquis, appearing natural but likely an escaped cultivar, Komunal
Budva, Petrovac, betw^een coasts of Lucica and Buljarica, forest rd. ca. 42° 12' N, 18° 57' E, 30 m, 24
Aug 2015, Coll. Tugrul Mataraci, 2015-28, Lab Acc: Robert P. Adams 14672,
cultivated, strict, columnar trees, in the park, Komunal Budva, Petrovac, Sv, Stefab coast, 42° 12' N,
18° 57' E, 2 m, 24 Aug 2015, Coll. Tugi'ul Mataraci, 2075-29, Lab Acc; Robert P. Adams 14673.
Turkey:
cultivated. Ayvalik- Town cemetery, Balikesir Province, living hedge around the cemetery, up to 20m
tall, strict habit, 39° 17' N, 26° 41' E, ca. 50 m, 18 July 2015, Coll. Tugrul Mataraci, 2015-24, Lab Acc:
Robert P. Adams 14597,
cultivated in park, Istanbul, Beyoglu-Halicioglu jet., strict habit, 41° 29' N, 28° 56' E , 34 m, 12 Aug
2015, Coll. Tii^iil Mataraci, 2015-25, Lab Acc: Robert P. Adams 14647,
cultivated on the highway between Izmit-Kocaeli, strict habit, 40° 46' N, 29° 39' E, 26m, 16 Aug
2015, Coll. Tugrul Mataraci, 2015-26, Lab Acc: Robert P. Adams 14648,
cultivated, Emirgan Park. Istanbul,strict, columnar tiees, 41° IF N, 29° 05' E, 84 m, 6 Sept 2015,
Coll. Tugrul Mataraci, 2015-30, Lab Acc: Robert P. Adams 14674,
50
Phytologia (Jan 19, 2017) 99(1)
United States:
cultivated. Carlsbad. CA, approx. 33° 06' 56.6" N, 117° 18' 39.3" W., 151 ft, 17 July 2015, San Diego
Co., Coll. Jim A. Bartel, 1631-1635. Lab Acc. Robert P. Adams 14591-14595. Dates trees 1631-1635
planted: 1985. 2005, 2000, 1980, 2010,
All specimens are deposited in the BAYLU herbarium.
Isolation of Oils - Fresh leaves (200 g) were steam distilled for 2 h using a circulatory Clevenger-
type apparatus (Adams, 1991). The oil samples were concentrated (ether trap removed) with nitrogen and
the samples stored at -20°C until analyzed. The extracted leaves were oven dried (100°C, 48 h) for
determination of oil yields.
Chemical Analyses - The oils were analyzed on a HP5971 MSD mass spectrometer, scan time 1
sec., directly coupled to a HP 5890 gas chromatograph, using a J & W DB-5, 0.26 mm x 30 m, 0.25
micron coating thiclcness, fused silica capillaiy colunm (see 5 for operating details). Identifications were
made by library searches of our volatile oil library (Adams, 2007), using tlie HP Chemstation library
search routines, coupled with retention time data of authentic reference compounds. Quantitation was by
FID on an HP 5890 gas chromatograph using a J & W DB-5, 0.26 mm x 30 m, 0.25 micron coating
thickness, fused silica capillary column using the HP Chemstation software.
RESULTS AND DISCUSSION
The volatile leaf oils of the horizontal form of C. sempenirens from natural populations in
Cyprus and Turkey were very unifonn and dominated (Table 1) by a-pinene (36.2, 26.0%), myrcene
(2.4, 2.4%), 8-3-carene (18.3, 16.0%), terpinolene (3.2, 3.8%), a-terpinyl acetate (4.7, 3.5%), cedrol (4.4,
3.3%), manoyl oxide (0.7, 3.8%), iso-pimara-7, 15-diene (0.4, 2.6%), isoabienol (2.4, 4.0%), and trans-
totai'ol (1.5, 5.7%).
The oils compositions of samples of C. sempen’irens cv. ‘Glauca Stricta’ from near San Diego,
CA, USA proved to very uniform, suggesting tliat these are likely clones. The average values of the
components show its oil to be quite similar to the horizontal form of C. sempervirens from natural
populations from Cyprus and Turkey (Table 1.) In contrast, the oils of the fastigiate foiins from Turkey
and Montenegro were quite variable (Table 1). Interestingly the oils from a cultivated tree and the 'wild'
(escaped cultivar?) fastigiate tree in Montenegro had quite different oils (Table 1).
Jacobson (1996) elaborated on the introduction and cultivation of Cupressus semperxurens
cultivars into the United States. He notes the introduction of the Italian cy press (cv. ‘Stricta’) into North
America is unknown, but George Washington planted one at Mt. Vernon in 1786. It seems vety^ probable
that Italitm cypress was introduced into Mexico by the Spaniards much earlier, as it is universally planted
at churches and cemeteries in Mexico. Jacobson (1996) lists the introductions of known cultivars as:
cv ‘Glauca Stricta’ < 1934; cv. ’Stricta’, date uncertain; cv. 'Swane’s Golden' <1977-78 by Swane Bros.
Nursery, Australia; cv. ‘Totem’ <1992, ex Duncan & Davies nursery, NZ; cv. ‘Variegata’ <1930s likely
from England ca. 1848. The commonly cultivated Italian cypress around San Diego, CA appears to be
cv. ‘Glauca Stricta.’
It is interesting that three components characteristic of cedamood oil (a-cedrene, p-cedrene,
cedrol) are present in the leaf oils from Cyprus, Turkey, ‘Stricta’ from California, and 14674 and 14647
from Turkey, but only a trace or absent from the other oils from fastigiate trees (Table 2). Overall, the
major teipenes compositions are very uniform for the horizontal fonn from natural populations (Cyprus,
Turkey) and cultivated fastigiate trees in California and Istanbul, but ver>^ variable (Table 2) for the other
cultivated fastigiate tree oils (Turkey and Montenegro).
Phytologia (Jan 19, 2017) 99(1)
51
a-pinene varies from 19.9% to 65.7% among the stricta oils (Table 2). In fact, the stricta Turkey
14597 is most unusual in having a high concentration of a-pinene, but very low concentrations of 5-3-
carene (0.2%), linalool (trace), a-cedrene (none), p-cedrene (0.1%), cedrol (trace) and abietadiene (trace).
Cultivated fastigiate Ciipressiis sempennrens trees from California and Istanbul (14674) both
have oils that are ver} similar to from natural populations from Cyprus and Turkey (Tables 1, 2).
Variation in tlie composition of oils from cultivated trees in Turkey and Montenegro suggests that these
cultivars arose from multiple selections of fastigiate trees, rather than cloning and subsequent widespread
cultivation. The volatile leaf oil composition does not support the recognition of the two growth forms of
C. sempervirem as distinct taxa. Similarly Faijon (2010) considered that the cultivated fastigiate form
was not a taxonomic variety but a cultigen.
ACKNOWLEDGEMENTS
Thanks to Amy TeBeest for lab assistance. This research was supported in part with funds from
Baylor University.
LITERATURE CITED
Adams, R. P. 1991. Cedarwood oil - Analysis and properties, pp. 159-173. in: Modem Methods of Plant
Analysis, New^ Series: Oil and Waxes. H.-F. Linskens and J. F. Jackson, eds. Springier- Verlag,
Berlin.
Adaius, R. P. 2007. Identification of essential oil components by gas chromatography/ mass
spectrometry. 2nd ed. Allured Pubf, Carol Stream, IL.
Adams, R. P., T, A. Zanoni, A. L. Cambil, A. F. Barrero, and L. G. Cool. 1997, Comparisons among
Cupressus arizomca Greene, C. benthamii Endl., C. lindleyi Klotz. ex Endl. and C. lusitanica Mill,
using leaf essential oils and DNA fmgeiprinting. J. Essential Oil Res. 9: 303-310.
Amri, I., L. Hamrouni, M, Hanana. S. Gargouri, and B. Jamoussi. 2013. Chemical composition, bio-
herbicidal and antifringal activities of essential oils isolated from Tunisian common cypress
{Cupressus sempennrens L.). J. Med. Plants Res. 7: 1070-1080.
Chanegriha, N., A. Baaliouamer, and B-Y. Meklati. 1997. GC and GC/MS leaf oil analysis of four
Algerian cypress species. J. Ess. Oil Res. 9: 555-559.
Farjon A. 2005. A monograph of Cupressaceae and Sciadopitys. Royal Botanic Gardens Press, Kew,
UK. 643 pp.
Farjon A. 2010. A handbook of the world's conifers. Brill Academic Publishers, Leiden, The
Netherlands. 1111 pp.
Floreani, S. A., J. A. Retamar, J. A. Retamar and E. G. Gros. 1981 . Essential oil of Cupressus
sempet'virens (cultivar ‘Stricta’). Essenze, Derivati Agmmari 51: 10-19.
Floreani, S. A., J. A. Retamar, and E. G. Gros. 1982. An. Asoc. Quimica Argentina 70: 663-667.
Gamero, J. P. Buil, D. Joulain, and R. Tabacchi. 1978. Parfums, Cosmetiques, Aromes 20: 33-36, 39-41.
Jacobson, A.L. 1996. North American landscape trees. Ten Speed Press, Berkeley, CA. 722 pp.
More, D, and J. White 2002 The illustrated encyclopedia of trees. Timber Press, Portland, OR. 800 pp.
Pauly, G., A. Yani, L. Piovetti, and C. Bernard-Dagan. 1983. Volatile constituents of the leaves of
Cupressus dupreziana and Cupressus sempeiMrens. Phytochemistry 22: 957-959.
S^kiewicz, K., K. Boratyhska, M. B. Dagher-Khamat, T. Ok, and A. Boratyhska. 2016. Taxonomic
differentiation of Cupressus sempenirens and C. atkmtica. Syst. and Biodiv. 14: 494-508.
Ulukanli, Z., S. Karaborklu, B. Ates, E. Erdogan, M. Cenet, and M. G. Karaastan. 2014. Chemical
composition of the esseniial oil fi'om Cupressus sempervirens L. horizontalis resin in conjunction
with it biological assessment. J. Ess. Oil-Bearmg Plants 17: 277-287.
52
Phytologia (Jan 19, 2017) 99(1)
Table 1. Leaf essential oil compositions for Cupressus sempervirens. Compounds in bold show large
differences between samples. Table abbreviations; horiz. = horizontal form, fast. = fastigiate Ibmi, Turk.
= Turkey, Calif = California, Tstan. = Istanbul, Mont. = Montenegro, Cyprus 15030 is the average of 5
samples (14560-14564); Turkey 15031 is the average of 4 samples (14564-14568); California is the
average of 5 samples (14591-14595). In these tliree cases, because little variation existed among the
samples, average oils are presented. All the other samples (Table 1) were collected from individual trees.
Mont, c 14673 is from a cultivated tree in Montenegro, whereas, Mont, ec 14672 is from an escaped
cultivar (?) tree in Montenegro.
KI
compound
horiz.
Cyprus
15030
m
fast.
Cahf
15032
fast.
Turk.
14647
fast.
Turk.
14648
fast.
Turk,
14597
fast.
Mont, c
14673
fast.
Mont, ec
14672
921
tricyclene
0.01
0.1
0.1
0 1
t
0.1
0,1
t
0.1
924
a-tliujene
0.02
0.1
0,1
0.5
0.1
t
t
1.8
932
a-pinene
36.2
26.0
39.1
34.4
28.5
35.2
65.7
19.9
29.6
945
a-fenchene
0.6
0.4
0.6
0.6
0.8
0.9
0.1
0.9
0.3
946
camphene
0.2
0.2
0,2
0,2
0.2
0.2
0.3
0.1
0.2
969
sabinene
0.5
0.6
0.7
3,4
0.4
0.4
1.0
1.2
3.6
974
(5-pinene
1.2
1.1
1.1
1.1
1.1
0.9
1.9
1.3
1.4
988
rayrccnc
2.4
2.4
2.2
2.4
2.3
2.3
2.6
2.7
3.9
1002
a-phel!andrene
t
t
t
t
t
t
t
t
t
1008
6-3-carene
18.3
16.0
16.8
17.3
30.1
25.7
0.2
30.7
12.2
1014
a-terpinene
0.2
0.2
0 1
0.2
0.1
0.1
t
0.2
0.5
1020
0,2
0.2
t
0.1
0.1
t
t
t
0.5
1023
sylvestrene
0.2
0.2
0.2
0.2
0,3
0.3
t
t
1024
1 imoncne
1.4
12
2.2
1.0
1.0
0.8
1.4
1.7
2.3
1025
P'phdlandrene
0.9
1,2
1.5
0.9
1.0
0.7
1.3
1.7
2.4
1044
(E)-|t-ocimene
t
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0.1
1054
y-terpinene
0.4
0.3
0,3
0,4
0.2
0,3
0,2
0.4
1.3
1067
linalool oxide
t
0.1
t
t
t
t
t
t
0.1
1082
m-cymenene
t
t
t
t
t
t
t
t
t
1086
terpiiiolcne
3.2
3.8
4.1
3.2
3.2
4.4
1.4
4.8
1.3
1099
linalool
1.5
0.6
0.2
0.3
0.6
t
t
0.4
1.1
1122
methyl octanoate
t
t
t
t
t
0.1
t
t
t
1123
a-camphenal
0.3
0.1
t
t
0.1
t
t
t
0,1
1133
cis-p-mentha-2,8-dien-l-ol
0.2
t
t
t
0.1
0.1
t
t
t
1135
trans-pinocar\’eol
0.2
t
t
t
t
0.1
t
t
t
1141
camphor
0,2
t
t
t
0.1
0.1
t
t
t
1154
karahanaenone
0,8
0.1
t
t
t
t
t
t
t
1159
p-mentha- 1 ,5-diene-8-ol,
isomer
0.3
0.1
t
t
0.2
0.1
t
t
t
1160
pinocarvone
0.2
t
t
t
t
t
t
t
t
1166
p-mentlia- 1 .5-diene-8-ol
t
t
t
t
t
t
t
t
t
1067
umbel lulone
0.1
0.3
t
t
t
t
t
t
t
1174
terpinen-4-ol
1.6
1.3
0.7
0.6
0.6
0.3
0,2
0.5
1.4
1176
m-c> men-8-ol
0.1
0.5
t
0.2
0.1
t
0.2
0.2
0.2
1179
p-c\ men-8-oI
0.2
0,1
t
t
0,1
t
t
t
t
1186
K-terpineol
0.3
0.2
0,2
t
0.2
0.1
t
0,1
0.1
12C4
myrtenol
t
0.2
t
t
t
t
t
t
t
1204
verbenone
0.3
0.1
t
t
0.2
0.1
t
t
t
1241
carvacrol. methyl ether
t
0,2
0.1
0.1
0.2
1.0
t
0.4
0.5
1254
linalool acetate
t
t
t
t
t
t
t
t
t
1287
bornyl acetate
0.2
0.2
0.1
0.4
0,2
0.1
t
0.8
1.5
1315
<2E,4E->decadi'enal
t
0.5
0.1
t
t
t
t
t
t
1323
meth\ l decanoate
t
t
t
t
t
t
t
t
t
1334
linalool propionate
0.6
0.7
0.4
0.4
1.3
0.7
t
1.0
0.4
1346
tt-terpiiiyl acetate
4.7
3.5
2.0
1.5
4.4
2.8
1.1
2.7
2.4
1345
«-cubebene
t
t
t
t
t
t
t
t
t
1374
a-ylangene
t
t
t
t
t
t
t
t
t
1400
tetradecane
t
0.1
t
t
t
0.1
t
t
0.1
1410
a-cedrenc
0.3
0.1
0.1
0.1
t
-
-
t
-
1411
2-epi-p-funebrene
t
0.1
0.1
0.1
t
-
-
t
-
1417
(E)-caiy ophyllene
0.1
0.2
0.1
0.2
0.3
t
0.1
0.4
0.8
1419
p-cedrene
0.3
0.3
0.1
0.2
0.3
t
0.1
0.4
t
Phytologia (Jan 19, 2017) 99(1)
53
KI
compound
horiz.
Cyprus
15030
horiz.
Turk,
15031
fast.
Calif
15032
fast.
Istan
14674
fast.
Turk.
14647
fast.
Turk.
14648
fast.
Turk
14597
fast.
Mont c
14673
fast.
Mont w
14672
1448
cis-muurola-3, 5-diene
0.3
0.3
0.1
0.1
t
0.6
0.2
0.4
0.3
1452
a-humulene
0.2
0.5
0.1
0.2
0.3
t
0.2
0.3
0.4
1465
cis-muurola-4l 14),5-diene
0,8
0.7
0,2
0.3
0.2
1.5
0.5
0.8
0.9
1478
y-muurolene
0.2
0.1
t
t
0.1
t
0.2
0.2
0.5
1480
^;ermacrene D
2.1
2.6
0.7
4.1
1.2
0.6
3.5
3.4
3.4
1499
epi-zonarene
0.2
0.2
t
t
t
0.6
t
0.2
0.3
1500
fjt-muurolene
0.1
0.1
t
t
t
t
0,3
0.1
0.1
1513
y-cadinene
0.1
t
t
t
t
t
t
0,1
0.2
1521
trans-calamenene
0,3
0.2
t
0.1
0.1
0.3
0.2
0.2
0.4
1.522
d-cadinene
0.3
0.2
t
0.1
0,2
0.2
0.2
0.2
0.3
i6mi
redrol
4,4
3.1
4.5
6.2
1.6
-
t
t
0.1
1652
a-cadinol
0.6
0.7
0.2
0.4
0.7
1.3
1.3
0.8
1.0
1685
germacra-4< 1 5).5, 1 0( 1 4 )-
trien-I-al
0.2
0.3
0.1
0.1
1.0
0.3
0.3
0.4
0.5
1921
meth’. 1 hexadecanoate
0.2
0.2
t
t
t
t
t
t
0.1
1958
iso-pimara-8( 1 4 ), 1 5-d iene
0.5
0.7
0.5
0.4
0.6
1.2
0.4
1.5
0.8
1987
mano> 1 o.xide
0.7
3.8
8.5
0.2
1.3
0.7
2.0
1.6
2.2
1987
iso-pimara-7, 15-diene
0.4
2.6
1.7
0.2
1.4
0.4
1.3
1.5
1.5
2055
abietatriene
1.5
3.4
1.4
0.5
0.9
1.6
2.5
1.2
1.1
2087
abietadiene
0.6
t
0.1
3.0
t
5.4
t
4.2
t
2103
6-octadecanoic acid,
methyl ester
0.4
t
t
t
t
0.2
t
t
0,5
2105
isoabienol
2.4
4.0
1.7
1.4
0.9
0.9
1.2
2.2
4.7
2149
abienol
0.4
1,3
1.0
3.2
0.4
0.8
0.2
1.1
0.9
2269
sandaracopimarinol
-
0.2
0.1
0.2
t
0.2
t
t
0.1
2282
sempervirol
t
0.4
0.1
t
t
t
t
t
0.1
2314
tiaiis-totarol
1.5
5.7
3.1
5.5
1.9
1.4
0.8
3.8
4.2
2331
trans-ferruginol
0.2
0.7
0.4
0.7
0.4
0.2
t
0.5
0.6
KI = linear Kovats Index on DB-5 column. Compositional values less than 0.1% are
denoted as traces (t). Unidentified components less than 0.5% are not reported.
Table 2. Comparison of the leaf oil eompositions for the most variable eompounds among samples.
KI
compound
horiz.
Cyprus
15030
horiz.
Turk,
15031
fast.
Calif
15032
fast.
Istan.
14674
fast.
Turk.
14647
fast.
Turk,
14648
fast.
Turk,
14597
fast.
Mont c
14673
fast.
Mont w
14672
932
a-pinene
36.2
26.0
39.1
34.4
28.5
35.2
65.7
19.9
29.6
1008
d-3-carene
18.3
16.0
16.8
17.3
30.1
25.7
0.2
30.7
12.2
1086
terpinolene
3.2
3.8
4.1
3.2
3.2
4.4
1.4
4.8
1.3
1099
linalool
1.5
0.6
0.2
0.3
0.6
t
t
0.4
1.1
1410
a-cedrene
0.3
0.1
0.1
0.1
t
-
-
t
-
1419
P-cedrene
0.3
0.3
0.1
0.2
0.3
t
0.1
0.4
t
1600
cedrol
4.4
3.1
4.5
6.2
1,6
-
t
t
0.1
1987
manoyl oxide
0.7
3.8
5.2
0.2
1.3
0.7
2.0
1.6
2.2
1987
iso-pima ra-7, 1 5-dieiie
0.4
2.6
5.2
0.2
1.4
0.4
1.3
1.5
1.5
2087
abietadiene
0.6
t
0.1
3.0
t
5.4
t
4.2
t
2105
isoabienol
2.4
4.0
1.7
1.4
0.9
0.9
1.2
2.2
4.7
2314
trans-totarol
1.5
5.7
3.1
5.5
1.9
1.4
0.8
3.8
4.2
54
Phytologia (Jan 19, 2017) 99(1)
Survey of cotton {Gossypium sp.) for non-polar, extractable hydrocarbons for
use as petrochemicals and liquid fuels
Robert P. Adams and Amy K. TeBeest
Baylor-Gmver Lab, Baylor University, 1 12 Main Ave., Gruver, TX 79040
robert_Adams@baylor.edu
James Frelichowski, Lori L. Hinze and Richard G. Percy
USDA-ARS, PA, SPARC, Crop Germplasm Research, 2881 F&B Road,
College Station, TX 77845
and
Mauricio Ulloa and John Burke
USDA-ARS, PA, CSRL, Plant Stress and Germplasm Development Research,
3810 4th Street, Lubbock, TX 79415
ABSTRACT
An ontogenetic study of a commercial cotton cultivar (FiberMax 1320), grown dryland, revealed
that the dry weight (DW) of leaves reached a maximum at the 1st flower stage, and then declined as bolls
opened. However, % pentane soluble hydrocai'bon (HC) yield continued to increase throughout the
growing season (due to the decline of leaf DW), It seems likely that as the bolls mature and seed are
filled, carbohydrates from the leaves are catabolized and sugars are transported to the bolls for utilization.
Per plant HC yields increased from square bud stage to 1st flower, remained constant until 1st boll set,
then declined at 1st boll-opened stage. This seems to imply that most of the HC are not catabolized and
converted to useable metabolites for filling bolls and seeds. A survey of arid land cotton accessions,
grown under limited iirigation or similar to dryland at Lubbock, TX, found % HC yield ranged from a
low of 2.88% to highs of 5.78 and 5.54% . Per plant HC yields ranged from 0.017 to 0.043 g/ g leaf DW .
Correlation between % HC yield and avg. leaf DW was non-significant (-0.103). A survey of USDA
germplasm cotton accessions, grown with supplemental underground drip imgation to achieve best yields
germinated by imgation, thence grown dryland at College Station, TX, found % HC yields were very
high, with four accessions yielding 11.34, 12.32, 13.23 and 13.73%. Per plant HC yields varied from
0.023 to 0.172 g/ g leaf DW. Hopi had a high % HC yield (10.03%), but it was the lowest per plant yield
(0.023 g/ g leaf DW). In contrast, China 86-1 with the second highest % HC yield (13.23%) was the
highest per plant yield (0.172 g). The correlation between % HC yield and avg. leaf DW was non-
significant (0.092). Thus, as seen in the arid land accessions, it appears that one might breed for both %
HC yield and leaf DW in cotton. Published on-line www.phytologia.org Phytologia 99(1): 54-61 (Jan
19, 2017). ISSN 030319430.
KEY WORDS: Cotton, Gossypium sp., yields of pentane extractable leaf hydrocarbons, petrochemicals,
liquid fuels.
There is a revived interest in sustainable, renewable sources of petrochemicals and fuels from ai‘id
and semi-arid land crops with the uncertainty of sustained crude oil production in the world. Adams et al.
(1986) screened 614 taxa from the western US for their hexane soluble hydrocarbon (HC) and resin
(methanol soluble) yields. They found the highest HC yielding species were from arid and semi-arid
lands in the Asteraceae (11 species), Asclepiadaceae (1), Celastraceae (1), Clusiaceae (1) and
Euphorbiaceae (1). The top 2% (12/614) had whole plant HC yields ranging from 10.4 to 16.4%.
Phytologia (Jan 19, 2017) 99(1)
55
Recently, Adams et al. (2017) surveyed native and cultivated sunflowers for their yields of leaf HC for
use as a potential semi-arid land crop and found high yielding (pentane extractable HC) plants. The top
2% had HC yields (ex leaves) ranging from
10.9 to 12.6% (Fig. 1), with the top 5%
ranging from 8.7% to 12.6%.
A preliminary analysis of the leaf HC
yields from six locally cultivated cotton
plants found a HC yield of 7.94% in one
plant. In comparison, HC yields from our
locally cultivated commercial sunflowers
ranged from 2.75 to 3.85%, as we expected,
in a crop that has been extensively selected
for seed production that leads to an
inadvertent selection against the production of
protective phytochemicals in the leaves.
A comparison between sunflowers and cotton characteristics shows considerable differences:
Figure 1 . Frequency distribution of HC yields for
329 H. annuus plants (from Adams et al., 2017).
characteristics
sunflowers (commercial)
cotton (commercial)
annual/ perennial
annual
perennial (but grown as an annual)
habit
herbaceous
woody
flowering
natural, 1 flower head/plant
(natural: many heads/plant)
induced by growth regulators or drought,
many flowers/plant (depending on
photoperiod)
leaf life
lower leaves yellow and die
generally defoliate for harvest
natural habitat
temperate. North America
dry tropics (or dry sub-tropics)
origin
from H. annuus. North America
complex genetics from taxa from around the
world (Wendel and Grover, 2015).
Annual sunflowers, herbaceous plants, live only to reproduce (by seed), whereas cotton, a woody,
perennial, having evolved with a dry season to induce flowering and seed formation, has adapted to a long
lifetime, in which annual seed production is not critical for short-term survival. However, maintaining
plant defensive chemicals and storing energy metabolites for cotton to survive the dry season are
important.
The evolution of modern cotton (Gossypium sp.) encompasses an improbable series of events that
involved transoceanic, long-distance dispersal with hybridization involving two diploids, one from the
Old World and one from the New World, forming the modern cultivated allo-tetraploid, G. hirsutum (the
reader is urged to read the informative account by Wendel and Grover, 2015).
Although there are several papers on the conversion of cotton field stubble to liquid fuels (see
Putun, 2010; Putuan et al., 2006; Akhtar and Amin, 2011 and references therein), there appear to be no
surveys of the yields of non-polar HC extractables in cotton.
As a part of our research on the investigation of contemporary crops for alternative, renewable
sources of petrochemical feedstocks and fuels, the present paper reports on the yields of HC from cotton
(Gossypium sp.) cultivars and accessions.
56
Phytologia (Jan 19, 2017) 99(1)
MATERIALS AND METHODS
Plant Materials:
Ontogenetic variation in HC yields study:
Commercial, cultivated cotton - FiberMax 1320, dryland, dark, loam soil, JP TeBeest Farm, 36° 25' 0.6"
N, 101° 32' 17.3" W, 3258 ft., Oslo, TX, avg. annual rainfall, 19.3". The eight (8) lowest growing, non-
yellowed mature leaves were collected at random from each of 10 cotton plants, at square bud, 1st open
flower, 1st boll, and 1st boll completely opened stages. The leaves were ah dried in paper bags at 49° C
in a plant dryer for 24 hr or until 7% moisture was attained.
HC yields of 30 cotton accessions representing photoperiodic and non-photoperiodic fonns of two
species:
Cultivated at the USDA-ARS Southern Plains Agricultural Research Center, College Station, TX, 30 37'
5.00" N, 96 21' 50" W, 354 ft., subsurface drip irrigation, sandy soil, annual rainfall 40". The lowest
growing, non-yellowed, mature leaf was collected at random, from each of 4-5cotton plants and bulked
for an accession sample. Different accessions varied in growth stage from square bud, 1st flower, and 1st
boll as the accessions were being grown for seed production. These accessions represent both
photoperiodic and non-photoperiodic types as well as obsolete cultivars within the two commercial
tetraploid cotton species, G. hirsutum and G. barbadense. These accessions were collected worldwide
and are maintained by the USDA National Cotton Germplasm Collection.
HC yields 21 cotton accessions grown for drought testing:
Cultivated at the USDA-ARS Plant Stress and Germplasm Development Research Center, Lubbock, TX,
33 35' 36.3" N, 101 54' 4.2" W, 3243 ft., hght, sandy soil, avg. annual rainfall 19.2". The lowest growing,
non-yellowed, mature leaf was collected at random, from each of 10 cotton plants and bulked for an
accession sample. Different accessions varied in growth stage from square bud to 1st flower. Some
supplemental water was applied during the growing season to attain germination and 1 united growth to
reflect plant stress responses, similar to dryland production, otherwise the plants were watered only by
natural rainfall. These accessions represent a diverse pool of G. hirsutum germplasm with different
genetic backgrounds from the USDA National Cotton Germplasm Collection.
Leaves were ground in a coffee mill (Imm). 3 g of air dried material (7% moisture) was placed
in a 125 ml, screw cap jar with 20 ml pentane, the jar sealed, then placed on an orbital shaker for 18 hr.
The pentane soluble extract was decanted through a Whatman paper filter into a pre-weighed aluminum
pan and the pentane evaporated on a hot plate (50°C) in a hood. The pan with hydrocarbon extract was
weighed and tared.
The shaker-pentane extracted HC yields are not as efficient as soxhlet extraction, but much faster
to accomplish. To correct the pentane yields to soxhlet yields, one sample was extracted in triplicate by
soxhlet with pentane for 8 hrs. All shaker extraction yields were conected to oven dry wt. (ODW)
multiplication of 1.085. For the cultivated TeBeost cotton, the shaker yields were corrected by the
increased soxhlet extraction efficiency (CF = xL56). For the arid land accessions, the soxhlet CF was
xl.31 and for the accessions grown at College Station, the soxhlet CF was xl.69.
RESULTS
Ontogenetic variation in HC yields in FiberMax 1320, grown dryland, are given in Table 1.
Notice (Fig. 2) that the D W of 8 leaves (Ivs) (from each plant) reach a maximum at the 1 st flower stage,
and then declined. However, % HC yield continued to increase throughout the growing season (due to
the decline of leaf DW). It seems likely that as the bolls mature and seed are filled, carbohydrates from
the leaves are metabolized into sugars that are transported to the bolls for utilization. Non-polar
hydrocarbons such as waxes, terpene hydrocarbons, alkanes, alkenes, etc. are thought to be largely inert
Phytologia (Jan 19, 2017) 99(1)
57
and not subject to catabolism. Notice that non-polar hydrocarbons (HC, as g DW/ 8 leaves) increased
from square bud stage to 1st flower, remained constant until 1st boll-set, then declined at 1st boll-opened
stage (Fig. 2). This seems to imply that most (-80% 0.355 g/0.440 g. Table 1) of the HC are not
catabolized and converted to sugars or other metabolites that might be utilized for during the maturation
of the bolls and seeds. Approximately -80% of the non-polar hydrocarbons remain in the leaves (at least
through the boll-opening stage (additional research is in planned to further examine the fate of non-polar
HC).
Table 1. Ontogenetic variation in pentane soluble hydrocarbon (HC) yields in FiberMax 1320, grown
dryland using eight leaves (Ivs) per plants and dry weight (DW) of leaves.
collection growth stage
DW for
8 Ivs/plant,
std err. mean
% HC yield,
std err. mean
Range of
yields(%)
HC g/ 8 Ivs DW,
std err. mean
14949 Cotton, cult Oslo,
square bud stage
5.49 g, 0.32
4.05%, 0.15
(3.31 - 4.56)
0.222 g, 0.016
14949 Cotton, cult Oslo,
1 St flower stage
7.46 g, 0.34
6.05%, 0.35
(4.78 - 7.84)
0.451 g, 0.053
14949 Cotton, cult Oslo,
1st boll set
6.29 g, 0.36
6.99%, 0.31
(4.95 - 8.28)
0.440 g, 0.034
14949 Cotton, cult Oslo, 1st
boll open, seeds maturing
4.43 g, 0.286
8.02%, 0.25
(6.65 - 8.90)
0.355 g, 0.027
Figure 2. Ontogenetic variation in HC yields (as % HC yield and g HC/g dry leaves) in FiberMax 1320.
The survey of arid land cotton accessions growing dryland at Lubbock, TX revealed (Table 2)
that % HC yield ranged from a low of 2.88% (14972, 16TXLWSA057) to highs of 5.78% (14961,
58
Phytologia (Jan 19, 2017) 99(1)
16TXLWSA039) and 5.54% (14964, 16TXLWSA043). Yields based on g HC/ g leaf DW ranged from
0.017 (14971, 16TXLWSA056) to 0.043 (14961, 16TXLWSA039 and 14965, 16TXLWSA047).
The con'elation between % HC yield and avg, leaf DW was non-significant (r = -0.103). Thus,
one might be able to breed for increases (up to some point) in both % HC yield and leaf DW in the same
genotype.
Table 2. Cotton screening for leaf HC of arid land accessions at USDA, Lubbock, TX.
Lab # , Plot No.
USDA
identifier
g avg leaf
DW
% yield HC*
g HC yield/ g
leafDW
14954 LI, 16TXLWSA002
DP1212
0.579
4.87
0.028
14955 L2 , 16TXLWSA012
SA-0464
0.542
5.30
0.029
14956 L3 , 16TXLWSA015
SA-0476
0.733
4.54
0.033
14957 L4 , 16TXLWSA016
SA-1049
0.600
3.83
0.023
14958 L5 , 16TXLWSA021
SA-1598
0.530
4.83
0.026
14959 L6 , 16TXLWSA029
STV5458
0.570
4.35
0.025
14960 L7 , 16TXLWSA036
SA-0473
0.493
3.69
0.018
14961 L8 , 16TXLWSA039
SA-1484
0.737
5.78 Hi 1
0.043 Hi
14962 L9 , 16TXLWSA041
SA-1269
0.627
4.08
0.027
14963 LIO , 16TXLWSA042
SA-1555
0,635
3.55
0.023
14964 LI 1 , 16TXLWSA043
SA-3128
0.632
5.54 Hi 2
0.035
14965 L12 , 16TXLWSA047
SA-2289
0.852
5.06
0.043 Hi
14966 L13 , 16TXLWSA049
FM2011
0.781
3.31 Lo
0.026
14967 L14 , 16TXLWSA050
PHY72
0.627
4.20
0.026
14968 L15 , 16TXLWSA052
SA-1762
0.836
3.49
0.029
14969 L16 , 16TXLWSA053
SA- 1 759
0.647
3.73
0.024
14970 LI 7 , 16TXLWSA055
SA-0429
0.682
3.88
0.026
14971 L18 , 16TXLWSA056
STV474
0.471
3.55
0.017 Lo
14972 LI 9 , 16TXLWSA057
PHY375
0.917
2.88 Lo
0.026
14973 L20 , 16TXLWSA059
SA-2169
0.773
4.59
0.035
14974 L21 , 16TXLWSA062
SA-1599
0.893
4.34
0.039
14975 L22 , Pima,
SJ-FR05
1.018
2.78
0.028
r (leaf wt, % yield) = -0.103 ns
The survey of USDA germplasm cotton accessions grown with supplemental uxigation at College
Station, TX, found % HC yields were very high, with four accessions yielding 1 1.34, 12.32, 13.23 and
13.73% (Table 3). These HC yields are in the top 2% reported by Adams et al. (1986) and top 1% for
sunflowers (Fig. 1, Adams et al. 2017).
Per plant HC yields (g HC/ g leaf DW) varied from 0.023 g to 0. 1 72 g, a 7-fold range (Table 3).
Hopi (14992) had a high % HC yield (10.03%), but it was the lowest per plant HC yield (0.023 g/ plant).
In contrast, China 86-1 (14997) with the second highest % HC yield (13.23%), had the highest per plant
HC yield (0.172, Table 3). The correlation between % HC yield and avg. leaf DW was non-significant (r
= 0.092 ns). Thus, as seen in the arid land accessions, it appears that one might breed (up to some
maximum point) for both % HC yield and leaf DW in cotton. This seems counter intuitive, but it may be
that cotton, being a perennial, and closely related to wild plants, may use the leaf hydrocarbons for plant
defensive chemicals. If so, there may be an evolutionary advantage to fully protect plants with large
leaves as well as those with small leaves. At this survey stage, we have not examined the amount of
gossypol (a known defense chemical).
Phytologia (Jan 19, 2017) 99(1)
59
Table 3. Cotton screening for leaf HC at USDA germplasm center, College Station, TX.
For % yield HC: + = 10.01 - 11.00%; ++ = 11.01 - 13.73%.
For g HC yield/leaf DW: + = 0.110 - 0.137g (top 13%); ++ = 0138 - 0.172g (top 3%).
Lab acc Source
USDA
identifier
g avg leaf
DW (# plants)
% yield HC
g HC yield/ g
leaf D W
14983, U1 , Tanguisw LMW 12-40
GB-0085
1 .335 (4)
5.97
0.080
14984, U2, Mono 57
GB-0204
1.360(4)
7.37
0.100
14985, U3, Nevis 81
GB-0227
0.728 (4)
10.36 +
0.041
14986, U4, Ashmouni Giza 32
' GB-0230
1.128(4)
7.37
0.083
14987, U5,Ashabad 1615
GB-0790
0.866 (4)
7.01
0.061
14988, U6, Tadla 2
GB-1439
1.106(4) !
9.70
0.107
14989, U7, 3-79
na
0.720 (4) ;
7.06
0.051
14990, U8, Pima S-5
S A- 1497
0.995 (4) '
‘ 7.92
0.079
14991, U9, TAM 87N-5
SA-1710
0.764 (4)
6.64
^ 0.051
14992, U 10, Hopi
^ SA-0033
1 0.266 (4)
10.03 +
0.023 Low
14993, Ull, Mexican #68
SA-0815
' 0.994 (4)
7.92
0.079
14994,U12,Christidis 53D7
SA-1 166
0.706 (4)
13.73 ++Hi
0.097
14995, U13,Acala SJ-1
SA-1 181
0.962 (4)
12.32 ++
0.119 +
14996, U 14, 3010
SA-1403
1.463 (4)
9.08
0.133 +
14997, U 15, China 86-1
SA-1419
1.300(4)
13.23 ++
0.172 ++Hi
14998, U 16, TM 1
SA-2269
1.244(4)
1 1 .09 ++
0.138 +
14999, U 17, KL 85/335
SA-2589
0.812(4)
10.25 +
0.083
15000, U18, KLM-2026
SA-2597
0.802 (4)
9.02
0.072
15001, U19,TAM91C-34
SA-2910
1.006 (4)
10.85 +
0.109
1 5002 ,U20, Vir-7080Col.Macias 1 7
SA-3348
0.896 (4)
11.34 ++
0.102
15003, U21,Palmeri, wild
TX-0005
: 0.398 (5)
7.92
0.032
, 15004, U22,Latifolium, wild
TX-OlOO
' 0.894 (5)
10.72 +
0.096
15005, U23,Latifolium, wild
TX-0104
0.967 (5)
9.25
0.089
15006, U24,Punctatum, wild
TX-0114
'0.815(5) ;
6.33
0.052
15007, U25,Morrili, wild
TX-0130
1 0.830 (5)
8.67
0.072
15008, U26,Marie-galante, wild
TX-0367
1.289(5)
7.37
0.095
15009, U27,Rlchmondi, wild
TX-0462
0.973 (5)
9.93
0.097
15010, U2S,Marie-galante, wild
TX-0866
0.511 (5)
8.05
0.041
15011, U29,Marie-galante, wild
TX-0878
0.692 (5)
4.50
i 0.031
15012, U30,Yucantanense, wild
TX-1046
0.728 (5)
3.29 Low
0.024 Low
r (leaf wt, % yield) = 0.092 ns
Principal Coordinate Analysis (PCoA), utilizing 597 SSR bands, of the 30 accessions revealed the
accessions are divided into G. barbadense and G. hirsutum (Fig. 3, left and right) (see Hinze et al., 2016
for further details on molecular marker analysis). The G. barbadense samples (8) are all improved
accessions. The samples of G. hirsutum contain both wild and improved accessions fonning a very loose
group, but the wild accessions are mostly found in the upper-right quadrant of the ordination (Fig. 3).
Utilizing the g HC/ g leaf DW data, the above average HC yielding accessions are clearly
clustered in a tightly grouped set of improved accessions (Fig. 3, dashed oval). Plotting the high and
highest yielding samples revealed that all three of the high yielding samples ( SA-1 181, SA-1403, SA-
2269, top 13%) and the highest yielding individual (SA-1419, top 3%) are found in that group (Fig. 3,
60
Phytologia (Jan 19, 2017) 99(1)
dashed oval). The discovery of the highest yielding individuals in a group of improved accessions is
surprising, in view of the selection for increased cotton seed and fiber yields.
Figure 3. Principal Coordinate Analysis (PCoA) based on 597 SSR bands. The percent of variance
accounted for among accessions is given on Dim 1 and Dim 2. See text for discussion.
It is also surprising that none of the wild accessions had high yields, although TX-OlOO had a
high % yield (10.72%), but having smaller leaves resulted in a moderate total g HC/ g leaf DW yield
(Table 3). It is interesting that genetically (by SSR data), TX-OlOO is ordinated nearest of any other wild
accessions to the high HC yielding group (Fig. 3). It may be that back-crossing TX-OlOO with S A- 1419
might produce some useful progeny in the future.
CONCLUSION
By the very definition of 'survey', this report is preliminary. Nevertheless, it seems remarkable
that a commercial crop, that has been bred and selected for seed (and lint) production, would sequester
such high amounts of hydrocarbons in leaves, as found in many cotton accessions. These results raise
many evolutionary questions, as well as numerous practical questions such as: Are the HC yields
heritable? Are they environmentally induced? Can breeding increase these HC levels without
detrimental effects on growth and hardiness? Clearly, much more research is needed (in progress).
LITERATURE CITED
Adams, R. P., M. F. Balandrin, K. J. Brown, G. A. Stone and S. M. Gruel. 1986. Extraction of liquid
fuels and chemical from terrestrial higher plants. Part I. Yields from a survey of 614 western United
States plant taxa. Biomass 9: 255-292.
Adams, R. P. and A. K. TeBeest. 2016. The effects of gibberellic acid (GA3), Ethrel, seed soaking and
pre-treatment storage temperatures on seed germination of Helianthus annuus and H. petiolaris.
Phytologia 98: 213-218.
Phytologia (Jan 19, 2017) 99(1)
61
Adams, R. P., A. K. TeBeest, B. Vaverka and C. Bensch. 2016. Ontogenetic variation in pentane
exti'actable hydrocarbons from HeUanthus annuus. Phytologia 98: 290-297.
Adams, R. P,, A. K. TeBeest, W. Holmes, J, A. Bartel, M, Corbet and D. Thornburg. 2017. Geographic
variation in pentane extractable hydrocarbons in natural populations of HeUanthus annuus
(Asteraceae, Sunflowers). Phytologia 99: 1-9.
Aklitar, J. and N. A. S. Amin. 201 1. A review on process conditions for optimum bio-oil yield in
hydrothermal liquefaction of biomass. Renewable and sustainable Energy Reviews 15: 1615-1624.
Hinze, L.L., E. Gazave, M.A. Gore, D.D. Fang, B.E. Scheffler, J.Z. Yu, D.C. Jones, J. Frelichowski and
R.G. Percy. 2016. Genetic diversity of the two commercial tetraploid cotton species in the
Gossypiurn Diversity Reference Set, Journal of Heredity 107: 274-286.
Putun, A. E. 2010. Biomass to bio-oil via fast pyrolysis of cotton sti'aw and stalk. J. Energy Sources 24:
275-285.
Putun, E., B. B. Urzun and A. E. Putun. 2006. Fixed-bed catalytic pyrolysis of cotton-seed cake: Effects
of pyrolysis temperatue, natural zeolite content and sweeping gas flow rate. Bioresource Technology
97: 701-701.
Wendel, J. F. and C. E. Grover. 2015. Taxonomy and evolution of the cotton genus, Gossypiurn. In:
Cotton, 2nd ed., D. D. Fang and R. G. Percy, eds.. Agronomy Monograph 57.
62
Phytologia (Jan 19, 2017) 99(1)
DNA sequencing and taxonomy of unusual serrate Juniperus from Mexico; Chloroplast capture
and incomplete lineage sorting in J. coahuilensis and allied taxa
Robert P. Adams
Biology Department, Baylor University, Box 97388, Waco, TX 76798, USA, Robert_Adams@baylor.edu
M. Socorro Gonzalez-Elizondo, Martha Gonzalez-Elizondo, David Ramirez Noya
CIIDIR Unidad Durango, Instituto Politecnico Nacional,
Sigma 119, Durango, Dgo., 34234 Mexico
and
Andrea E. Schwarzbach
Department of Health and Biomedical Sciences, University of Texas - Rio Grande Valley,
Brownsville, TX 78520, USA.
ABSTRACT
Analysis of nrDNA, petN-psbM, trnS-trnG, trmD-trnT, and tmF-trnL of Juniperus coahuilensis
and allied taxa of Mexico found typical J. coahuilensis, as well as individuals with: coahuilensis cp and
hybrid ITS; coahuilensis cp and novel ITS sequence (La Pamlla type); novel Blue Fruited cp (blue fruited
taxon) and coahuilensis ITS; plus Blue Fruited cp and La Parrila ITS. nrDNA data was examined and
found to detect hybridization, chloropla.st capture and incomplete lineage sorting. In addition, a new
taxon was found with Blue Fruited (Blue Fruited) cp and /. martinezii ITS, suggestive of chloroplast
capture. New records of J. saltillensis were confimied from Zacatecas. A new record of J. martinezii
from Durango was also confirmed. Several plants affiliated with eitlier J. martinezii, or J. flacckla were
in distinct clades showing the need for additional research on their volatile leaf oils, moiphology and
ecology to address their taxonomic status. And lastly, a very unusual population of junipers large, single
stemmed trees with aff. J. poblana was found in Nayarit, with long and pendulous foliage. Analysis of
the leaf volatile oils, ecology and morphology of this taxon is necessary (in progress) to ascertain its
taxonomic rank. Published on-line www.phytologia.org Phytologia 99(1): 62-73 (Jan 19, 2017). ISSN
030319430,
KEY WORDS: Juniperus coahuilensis, J.flaccida, J. martinezii, J. poblana, Cupressaceae,
hybridization, introgression, incomplete lineage sorting, nrDNA polymorphisms, petN-psbM DNA.
As a part of on-going research on Juniperus, recently, Adams (2016) found (by petN-psbM
sequencing) that Juniperus arizonica, previously known only from Arizona and New Mexico, occurs in
northern Sonora and Chihuahua, trans-Pecos Texas in the Franklin Mtns., Hueco Mtns., Hueco Tanks
State Park, Quitman Mtns., Eagle Mtns. and Sierra Vieja Mms., primarily on igneous material. These
trans-Pecos juniper populations have previously been identified as J, coahuilensis.
Additional examination of populations of J. coahuilensis in the Trans-Pecos, Texas region
(Adams 2017) revealed that situation was more complex with a relatively sharp demarcation between J.
arizonica and J. coahuilensis (Fig. 1). The zone of contact and likely hybridization is in Hueco Tanks
State Park, Quitman Mtns., and Anima Mtns. and this appears to be a region of introgression northward
from J. coahuilensis (Fig. 1).
Although it appeared that the J. coahuilensis at La Zarca, MX was a pure population (Adams
2017), new specimens of aff. J. coahuilensis with violet, reddish and blue colored fruits have been
discovered in north central Mexico that do not fit the current Juniperus keys (Adams 2014). The present
distribution map of J. coahuilensis is shown in Figure 2.
Phytologia (Jan 19, 2017) 99(1)
63
The purpose of this paper is to report on the results of DNA sequencing for these new,
morphologically variable samples in an effort to better understand the variation in the serrate junipers of
Mexico, with particular emphasis on J. coahuilensis and its allies.
Figure 1 . Plant distribution map
showing their classification as J.
arizonica, J. coahuilensis, or hybrids
based on the results from both nrDNA
and cpDNA analysis. From Adams
(2017).
MATERIALS AND METHODS
Plant material and populations studied:
J. coahuilensis, large population with thousands of trees. Mexico, Durango, 85 km n. of La Zarca on Mex.
45, 26° 21' N, 105° 16.66’ W,1740m, 10 Dec 1991, Robert P. Adams 6829-6831,
J. coahuilensis, large population in Bouteloua grassland, multi-stemmed tree, 4 m tall, female, female
cones glaucous, blue-pinkish when mature. Mexico, Durango, at km 18 on Mex. 45, north of Durango,
pollen shed in fall, bark exfoliating in narrow strips. 24° 09.067’N, 104° 42.462’ W, 1938 m, 7 May
2004, Coll. R. P. Adams 10241, 10242.
J. aff. coahuilensis, shrub or tree 3-6 m, seed cones globose, fleshy, bright rose to salmon colored, sweet,
l(2)-seeded, on limestone, Mexico, Durango, Mpio. Nombre de Dios, San Jose de La Parrilla, 23° 44'
20" N, 104° 07' 20" W, 2120 m, 27 Aug 2004, Coll. Socorro Gonzalez 6988, Lab Ace. Robert P. Adams
10454,
J. aff. coahuilensis. Plant on limestone, with unusual seed cones: fibrous, bluish appearance because of
the dense glaucous cover on a green surface, one seed [Not fleshy, nor rose or salmon, nor sweetish as
in J. coahuilensis]-, bark thin, fibrous, gray-brown, Mexico, Durango, Mpio. Nombre de Dios, San Jose
de La Parrilla; on limestone, 23° 44' 20" N, 104° 7' 20" W, 2120 m, 27 Aug 2004, Coll. Socorro
Gonzalez 6989, Lab Ace. Robert P. Adams 10455,
J. aff. coahuilensis hybrid?. Plant with unusual seed cones: fleshy as found in J. coahuilensis (present in
the same site), but differs having dull purple to dull rose color, glaucous, seed cones in dense groups;
branches firm, ascendant; bark thin, fibrous, gray-brown, Mexico, Durango, Mpio. Nombre de Dios,
San Jose de La Parrilla; on limestone. 23° 44' 20" N, 104° T 20" W, 2120 m, 27 Aug 2004, Coll.
Socorro Gonzalez 6990; Lab Ace. Robert P. Adams 10456,
J. aff. coahuilensis. Abundant shrubs, 2-3 m, seed cones rose-pale cherry, without glaucous cover,
Mexico, Durango, Mpio. Guanacevi; SE of Guanacevi, on road to Durango, 25° 53' 14" N, 105° 50' 59"
64
Phytologia (Jan 19, 2017) 99(1)
W, 1990 m, 27 Aug 2004, Coll. Socorro Gonzalez and M. Gonzalez-Elizondo7005; Lab Acc. Robert P.
Adams 10459,
J. aff. coahuilensis, Abundant, trees on limestone, to 3 m, seed cones fleshy, red-orange, sweet, Mexico,
Durango, Mpio. Nombre de Dios, S of El Porvenir and NE of San Jose de La Parrilla, 23° 46' 30" N,
104° 09’ 30" W, 1980 m, 4 Nov 2004, Coll. Socorro Gonzalez 7016-1, 7016-2, Lab Acc. Robert P.
Adams 10503, 10504,
J. aff. coahuilerms, shrub-trees, on limestone, seed cones violet colored, somewhat fibrous and resinous,
Mexico, Durango, Mpio. Nombre de Dios, NE of San Jose de La Parrilla. 23° 46' N, 104° 9' W, 1980
m, 4 Nov 2004, Coll. Socorro Gonzalez 7017a, Lab Acc. Robert P. Adams 10505,
J. aff. coahuilensis, shrub-trees, on limestone, seed cones: densely grouped, fleshy, sweet, reddish-orange,
1(2) seeds; thin, fibrous bark: on branches pale gray to whitish, Mexico, Durango, Mpio. Nombre de
Dios, 0.4 km SW of San Jose de La Parrilla; on limestone, 23° 44' 20" N, 104° T 20" W, 2120 m, 4
Nov2004, Coll. Socorro Gonzalez 7019-1, 7019-2, Lab Acc. Robert P. Adams 10511, 10512,
J. cf, Jlaccida, Short trees, 1 .5-3 m tall; bark on branches papery and exfoliating, inner bark smooth,
reddish; no seed cones, similar to 7, Jlaccida, but in a very dry habitat in the Chihuahuan desert region,
Mexico, Durango, Mpio. Lerdo, Sierra del Rosario, nearly atop the mountain, with Yucca and oak
scrub; on limestone, 25° 38’ 44” N, 103° 54’ 40” W, 2700 m, 8 Apr 2008, Coll. M. S. Gonzalez-
Elizondo et al. 7375 a,b; Lab Acc. Robert P. Adams 14616, 14617.
J. aff. rnartinezii/ durangensis, Shrub, seed cones orangish color and fibrous, with pinyon pine and oaks.
Mexico, Durango, Mpio. Panuco, Sierra de Gamon, N\V slopes, 24° 35’ N, 104° 16’ W, 2500 m, 4 June
2008, Coll. M. S. Gonzalez-Elizondo et al. 7391 a,b; Lab Acc. Robert P. Adams 14618, 14619,
J. aff. saltillensis, Abundant shrub 1-1.8 m, dai‘k blue seed cones, somewhat glaucous, Mexico, Zacatecas,
Sierra de Mazapil, Mpio. Concepckin del Oro, 24° 37’ 21” N, 101° 28’ 0^5” W, 2850-2900 m, 16 Oct
2009, Coll. M. S. Gonzalez-Elizondo and M. Gonzalez-Elizondo 7567,7568; Lab Acc. Robert P. Adams
14620, 14621
J. aff. poblana, uncommon young trees (saplings) 2 m, in oak woodland dominated by Querciis resinosa,
Mexico, Nayarit, Mpio. El Nayar, SW of Mesa del Nayar on road to Ruiz, Km 86.8; S of bridge of
arroyo del Fraile, E of El Maguey, 22° 10’ 08” N, 104° 43’ 5 1” W, 1 150 m, 19 Jan. 2016, Coll. M. S.
Gonzalez-Elizondo and M. Gonzalez-Elizondo 8381 with L. Lopez, A. Torres Soto; Lab Acc. Robert P.
Adams 14896
J. aff. poblana, large, single stemmed trees, foliage long and pendulous, abundant ti'ees, up to 25 m high,
on strongly rocky slope, forest of Jimiperus-Clusia with elements of mesophytic forest (Magnolia) and
tropical forest (Bursera, Opuntia, Pilosocereus purpusii) as well as Agave attemiata and Yucca
jaliscensis, Mexico, Nayarit, Mpio. El Nayar, SW of Mesa del Nayar on road to Ruiz; NE of El
Maguey, 22° 07’40” N, 104° 47’ 47” W, 1430 m, 19 Jan. 2016, Coll. M. S. Gonzalez-Elizondo and M.
Gonzalez-Elizondo 8379a,b,c,d, with L. Lopez, A. Torres Soto; Lab Acc. Robert P. Adams 14897 -1 4900,
J. maninezii, new record for Durango, Abundant tree with drooping branchlets, pale grayish-green foliage
with white resin marks, Mexico, Durango, Mpio. Vicente Guerrero, Sierra de Organos, near the border
of state of Zacatecas, northernmost known population of J. martinezii. The closest population is about
220 km to the SE [Aguascalientes, San Jose de Gracia (acc, Perez de la Rosa 1985) 23° 47’ 28” N, 103°
49’ 44” W, 2225 m, 21 Jan 2016, Coll. M. S. Gonzalez-Elizondo and M. Gonzalez-Elizondo) 8384; Lab
Acc. Robert P. Adams 14901,
J. aff. coahuilensis. Shrub, blue seed cones, Mexico, Durango, Mpio, Nombre de Dios, 4 km w of San
Jose de La Parrilla, 23° 43’ N, 104° 08' W, 2150 m, 25 Oct 1983, Coll. M. S, Gonzalez-Elizondo et al.
2776; Lab Acc. Robert P. Adams 14902,
J. aff. coahuilensis. Shrub, blue seed cones, Mexico, Durango, Mpio. Tepehuanes, SE edge of town, 25°
20' N, 105° 43' W, 1800 m, 10 Sep 1989, O. Bravo 288-, Lab Acc. Robert P. Adams 14903,
J. aff, coahuilensis. Shrub, blue seed cones, Mexico, Durango, Mpio. Santiago Papasquiaro, 9 km por el
camino a Los Altares, 25° 06' N, 105° 27' W, 1940 m, 30 July 1990, Coll. A. Benitez P. 1646; Lab Acc.
Roben P. Adams 14904,
Voucher specimens for new collections are deposited in the Herbarium, Baylor University (BAYLU).
Phytologia (Jan 19, 2017) 99(1)
65
One gram (fresh weight) of the foliage was placed in 20 g of activated silica gel and transported
to the lab, thence stored at -20° C until the DNA was extracted. DNA was extracted from juniper leaves
by use of a Qiagen mini-plant kit (Qiagen, Valencia, CA) as per manufacturer's instructions.
Amplifications were performed in 30 pi reactions using 6 ng of genomic DNA, 1.5 units Epi-
centre Fail-Safe Taq polymerase, 15 pi 2x buffer E (petN-psbM), D (maldehy) or K (nrDNA) (final
concentration: 50 mM KCl, 50 mM Tris-HCl (pH 8.3), 200 pM each dNTP, plus Epi-Centre proprietary
enhancers with 1.5 - 3.5 mM MgCb according to the buffer used) 1.8 pM each primer. See Adams, Bartel
and Price (2009) for the ITS and petN-psbM primers utilized. The primers for tmD-tmT, tmL-trnF and
trnS-trnG regions have been previously reported (Adams and Kauffmann, 2010). The PCR reaction was
subjected to purification by agarose gel electrophoresis. In each case, the band was excised and purified
using a Qiagen QIAquick gel extraction kit (Qiagen, Valencia, CA). The gel purified DNA band with the
appropriate sequencing primer was sent to McLab Inc. (San Francisco) for sequencing. Sequences for
both strands were edited and a consensus sequence was produced using Chromas, version 2.31
(Technelysium Pty Ltd.) or Sequencher v. 5 (genecodes.com). Sequence datasets were analyzed using
Geneious v. R7 (Biomatters. Available from http://www.geneious.com/) , the MAFFT alignment program.
Further analyses utilized the Bayesian analysis software Mr. Bayes v.3.1 (Ronquist and Huelsenbeck
2003). For phylogenetic analyses, appropriate nucleotide substitution models were selected using
Modeltest v3.7 (Posada and Crandall 1998) and Akaike's information criterion.
RESULTS AND DISCUSSION
Sequencing nrDNA, petN-psbM, trnS-trnG, trnD-trnT and trnL-trnF resulted in 4,351 bp of
concatenated sequence data. A Bayesian tree shows the placement of most of the samples collected as J.
aff. coahuilensis (10241, 10242, 10503, 10504, 10505) are in the clade with typical J. coahuilensis
(shaded box. Fig. 2). However, an adjacent clade (cross-hatched box. Fig. 2) contains two sub-clades:
blue seed cones plants (14902, 14903, 14904) and La Parrilla plants, with very variable seed cone colors
from violet to bluish to orange (14055, 10454, 10456, 10459, 10511).
Plants 14620, 14621, J. aff. saltillensis from Zacatecas, Sierra de Mazapil, Mpio. Concepcion del
Oro, are nested, loosely in a clade with J. saltillensis (Fig. 2). Additional research on the leaf volatile
oils, ecology and morphology (in progress) may prove these to be a new variety of J. saltillensis.
Sample 14901, collected as J. martinezii from Durango, Mpio. Vicente Guerrero, Sierra de
Organos, near the border of state of Zacatecas, is in a clade with J. martinezii (Fig. 2). This is the first
report of J. martinezii from Durango and is the northernmost known population of J. martinezii. The
closest population is about 220 km to the SE (Aguascalientes, San Jose de Gracia, Perez de la Rosa,
1985).
Two other collections (14618, 14619, shrubs, seed cones orangish color and fibrous, with pinyon
pine and oaks. Mexico, Durango, Mpio. Panuco, Sierra de Gamon) with affinities to both J. martinezii
and J. durangensis, were placed in a clade with J. martinezii and J. durangensis (Fig. 2). There is some
support for it being in a distinct clade (51%, Fig. 2), but additional research is needed on the leaf volatile
oils, ecology and morphology (in progress) to determine if this taxon is a new variety of J. martinezii or
perhaps a new species.
Plants 14616, 14617, collected as J. ddf. flaccida, were short trees, 1.5-3 m tall with the bark on
branches papery and exfoliating, and inner bark smooth, reddish. These samples were in a well supported
66
Phytologia (Jan 19, 2017) 99(1)
10231
100
virginiana, out-gfoup
10232
-7^
100
7635
7636
10931
10932
anzonica
monos perm a
angosturana
■10463^
typical
coahuilensis
10512
10241
10504
6829
6830
14902
blue fruited
'aff. coah'
Bayesian Tree
nrDNA, petN-psbM, trnS-trnG
trnD-trnT, trnF-trnL
4,351 bp
14901 martinezii Dgo.
5950
100 ' — 5951
6852 , .
6853
14616
martinezii
14617
flaccid a
aff. flaccida
monticola
100 ^- 6877
^°°-Jf/jaliscana
11926 poblana v. decurrens
100 ' ^11927
r 14896
14897 Nayar it trees
14898 aff. poblana
- 14899
— 14900
Figure 2. Bayesian tree of
serrate leaved Juniperus of
North America. Numbers
next to branch points are
posterior probabilities as
percents. Note the typical
J. coahuilensis (shaded
box) and the adjacent clade
(cross-hatched box). See
text for discussion.
Samples in boldface print
are new collections.
Samples in regular font are
the reference set of serrate
junipers.
clade with J. flaccida, but yet, quite distinct (Fig. 2). The site is in a very dry habitat in the Chihuahuan
desert region, Mexico, Durango, Mpio. Lerdo, Sierra del Rosario. No seed cones were found (April,
2008), so new collections with seed cones are needed. Clearly, additional research is needed on the leaf
Phytologia (Jan 19, 2017) 99(1)
67
volatile oils, ecology and morphology (in progress) to determine if this taxon might be a new variety of J.
flaccida.
And lastly, a very unusual population with aff. poblana, was found with large, single stemmed
trees, and foliage long and pendulous in Nayarit. Analysis of their DNA did place them (14986, 14897,
14898, 14899, 14900) in a large clade with J. poblana and J. p. var. decurrens (Fig. 2). However, they
are quite distinct and well supported as a separate clade. Analysis of the leaf volatile oils, ecology and
morphology (in progress) should be sufficient to determine if this taxon is a new species, or perhaps
another (new) variety of J. poblana.
A detailed examination of
variable nrDNA (ITS) sites of J.
coahuilensis aff. samples, as well as J.
coahuilensis from the Trans-Pecos,
Texas region is shown in Table 1.
Overall, J. coahuilensis and the aff.
samples from Mexico do not have as
many variable sites as found in the
Trans-Pecos region (see also Adams,
2017).
Mapping the classification of
individuals based on ITS and cp
(petN) data shows (Fig. 3) only four
samples in Durango that have both
ITS and cpDNA of J. coahuilensis (as
found in the Trans-Pecos, Texas area).
The CpDNA of the blue
fruited taxon (black filled circles. Fig.
3) was found in combination with
various types of ITS DNA in central
and southern (La Parrilla area)
Durango. The cpDNA of typical J.
coahuilensis was found in both
northern and southern Durango (Fig.
3). Two of the blue fruited samples
(black filled circle, open diamond.
Fig. 3) were found in central
Durango, and the third sample was
found in the La Parrilla
area. Two samples with La Parrilla
type ITS (LaPar, Table 1; black square.
Fig. 3) were found in the La Parrilla
area and are in northwestern Durango.
Distribution of
J. coahuilensis, hybrids
and other taxa based on
ITS and cpDNA
^ Alpine
*Chi
a
cp coah
ITS coah
X
Q Cp coah
y ITS hybrid
8 cp Blue fruit
ITS coah
S ep coah
ITS La Parrilla
# cp Blue fruit
■ ITS La Parrila
^ cp Blue fruit
O ITS J. martinezii
•A
La Parrilla
area
Figure 3. Map of J. coahuilensis and aff. samples by their
CpDNA (petN) (circles) and ITS DNA (squares). Data in the
Trans-Pecos, Texas area from Adams (2017.
Two samples, putatively hybrids based on their ITS, were found in the La Parrilla area (crossed
squares. Fig. 3). All six of the cpDNA/ ITS types were found in the La Parrilla area (Fig. 3). It may be
that other areas are equally as diverse, but additional sampling is needed to address this question.
68
Phytologia (Jan 19, 2017) 99(1)
Several of the nrDNA (ITS) sites display interesting geographic patterns. ITS site 191 (A,G,
A/G) has considerable variation in the Trans-Pecos, Texas region (Fig. 4) and continues into northern
Durango. However, no other A/G sites were found in central and southern (La Parrilla) Durango. This
may be the result of hybridization/ introgression from some juniper in the Alpine area.
One individual with site 191 (A) was found south of Alpine and another found in the La Parrilla
area of southern Durango.
Figure 4. Geographic variation in ITS
site 191. See text for discussion.
Phytologia (Jan 19, 2017) 99(1)
69
ITS site 196 featured the deletion of T in many samples ranging from the Alpine to central and
southern Durango (Fig. 5). All three BF (blue fruited) and the 'LaPar' ITS type samples had the 196
deletion (Table 1). Plants 10454, 10456, and 14903 appear to be hybrids. The deletion caused slippage
during sequencing, so all the sites downstream from 196 were polymorphic. To remedy this problem, a
new internal reverse primer was synthesized and used to reverse-sequence the immediate 700 bp past site
196 to obtain clean sequences from some plants. It is not known if this deletion is of contemporary or
ancient origin.
Figure 5. Geographic variation in ITS
site 196. See text for discussion.
70
Phytologia (Jan 19, 2017) 99(1)
Mapping ITS site 303 provided a novel pattern not seen in other ITS sites. The presence of C/T
polymorphisms for site 303 in the Trans Pecos area (Fig. 6) was not found in Mexico (nor in AZ, NM, see
Adams, 2017). This seems to imply that the event was modern and due to hybridization with some
unknown extant or extinct juniper in the Trans-Pecos area. Of interest to this study was the finding many
plants with either C or T, but no plants with C/T in Durango.
In addition, the three BF (blue fruited) plants each contained G at site 303 (Table 1) and are
shown (Fig. 6) with two in central Durango and one in the La Parrilla area. In addition, G (site 303) is
also found in J. martinezii (Table 1). This site, no doubt, supported the placing of the BF junipers in a
clade with J. martinezii in a NJ tree based on ITS sequences (data not shown), suggesting the BF taxon
has a nuclear affinity to J. martinezii. However, sequences from the four cp gene regions was
concatenated to nrDNA data in the construction of the Bayesian tree (Fig. 2), and this led to the
positioning of the BF taxon loosely in the J. coahuilensis clade (Fig. 2).
Figure 6. Geographic variation in ITS
site 303. See text for discussion.
Phytologia (Jan 19, 2017) 99(1)
71
Finally, examination of ITS site 1116 presents an interesting situation in that every case with C/T
at site 1116 (Table 1) has a deletion (del) at 196 (Table 1). Re-examination of the nrDNA sequence for
14814 revealed that the site 196 contains mostly T, there is a small (ca. 20% C peak). From 196 onward,
small peaks (ca. 20% high) are present in the sequence. The del at 196, the slippage of the sequence for
ca. 20% of the DNA strains perfectly explains the minor bases from 196 onward. This suggests that the
plant is of backcross origin and that incomplete lineage sorting has not yet removed the minority copies
that contain a del in 196). It should be noted that several samples (Table 1) have a del at 196 but have
either a clean C or T at 1 1 16.
The pattern seen for site 1116 (Fig. 7) suggests (as seen in Fig. 5) hybridization throughout the
range of J. coahuilensis from Alpine to southern Durango, with the presence of numerous plants with C
or T at site 1116.
Figure 7. Geographic variation in ITS
site 1116. See text for discussion.
72
Phytologia (Jan 19, 2017) 99(1)
ACKNOWLEDGEMENTS
This research was supported in part with funds from Baylor University. Thanks to Amy TeBeest
for lab assistance.
LITERATURE CITED
Adams, R. P. 2014. The junipers of the world: The genus Jiiniperus. 4th ed. Trafford Publ., Victoria, BC.
Adams, R. P. 2016. Junipenis arizonica (R. P. Adams) R. P. Adams, new to Texas. Phytologia 98:179-
185.
Adams, R. P. 2017. Multiple evidences of past evolution are hidden in nrDNA of Junipenis arizonica
and 7. coalmilensis populations in the trans-Pecos, Texas region. Phytologia 99: 39-48.
Adams, R. P. J. A. Bartel and R. A. Price. 2009. A new genus, Hesperocyparis, for the cypresses of the
new world. Phytologia 91: 160-185.
Adams, R. P. and J. R. Kistler. 1991. Hybridization between Junipenis erythrocarpa Cory and Jiiniperus
pinchotii Sudworth in the Chisos Mountains, Texas. Southwest. Natl. 36: 295-301.
Adams, R. P., M. Miller and C. Low. 2016. Inheritance of nrDNA in artificial hybrids of Hesperocyparis
arizonica x H. macrocaipa. Phytologia 98: 277-283.
Adams, R. P. and A. E, Schwarzbach. 2011. DNA barcoding a juniper: the case of the south Texas
Duval county juniper and serrate junipers of North America. Phytologia 93(1): 146-154.
Adams, R. P. and A. E. Schwarzbach. 2013. Taxonomy of the serrate Xo^dif Jiiniperus of North America:
Phylogenetic analyses using nrDNA and four cpDNA regions. Phytologia 95: 172-178.
Adams, R. P. and A. E. Schwarzbach. 2015. A new, flaccid, decurrent leaf variety of Junipenis poblana
from Mexico: J. poblana var. deciurens R. P. Adams. Phytologia 97: 152-163.
Perez de la Rosa. J.A. 1985. Una nueva especie de Juniperus de Mexico. Phytologia 57: 81-86.
Posada, D. and K. A. Crandall. 1998. MODELTEST: testing the model of DNA substitution.
Bioinformatics 14: 817-818.
Ronquist, F. and J. P. Huelsenbeck. 2003. MrBayes 3: Bayesian phylogenetic inference under mixed
models. Bioinformatics 19: 1572-1574.
Phytologia (Jan 19, 2017) 99(1)
73
Table 1. Vai'iable sites in ni'DNA for J. arizonica (ariz), J. coahuilensis (coah) and 'blue, violet, bluish' fruited (BF),
and La Parrilla type nrDNA (LaPar). del = deletion, mart = J. nuirtinezii nrDNA.
sample
petN
ITS
191
196
302
303
304
318
533
543
1116
1148
#poly
azin634Sedona 181 A/C
ariz
ariz
G
T
A
C
T
T
A
C/G
T
C
2
az 1 063 5S edona 68 1 A/C
ariz
ariz
G
T
A
c
T
T
A
C/G
T
C
2
azlll636Sedona
ariz
ariz
G
T
A
c
T
T
A
C/G
T
C
1
az 1 4908Cottotiwood
ariz
ariz
G
T
A
c
T
T
A
G
T
C
0
az 1 4909Cottoriwood
ariz
ariz
G
T
A
c
T
T
A
C/G
T
C
1
az 1 49 1 OCottonwood
ariz
ariz
G
T
A
c
T
T
A
C
T
C
0
az 1 49 1 ICottonwood
ariz
ariz
G
T
A
c
T
T
A
c
T
C
0
az 149 13 Cotton wood 121C/T
ariz
ariz
G
T
A
c
T
T
A
C/G
T
C
2
az7635RockHoundSP
ariz
aiiz
G
T
A
c
T
T
A
C
T
C
0
az7636RockHoundSP
ariz
ariz
G
T
A
c
T
T
A
C
T
C
0
az7637RockHoundSP
ariz
ariz
G
T
A
c
T
T
A
C/G
T
C
1
azl0630RockHSP
ariz
ariz
G
T
A
c
T
T
A
C
T
C
0
coa 1 4807sofAlpine
coah
coah
G
del
A/G
C/T
C/T
C/1
T
C
C/T
C/T
6
coal4808sofAlpine
coah
coah
G
T
A
c
T
C/T
T
C
T
C/T
2
coa 1 481 OsofAlpine
coah
coah
G
del
A
C/T
C/T
T
T
C
C/T
C
4
coa 14811 sof Alpine
coah
coah
A
del
A/G
C/T
T
T
T
C
C/T
c
3
coa 1 48 1 2\vof Alpine
coah
coah
G
del
A/G
C/T
T
C/T
T
C
C
c
4
coal4813wofAlpine 313A/G
coah
coah
G
T
A
C/T
T
C/T
T
C
T
C/T
4
coa 1481 4wof Alpine
coah
coali
G
def^
A
C
T
T
T
C
C/T
C
1
coa 148 1 5\vof Alpine
coah
coah
G
del
A/G
C/T
T
C/T
T
C
C/T
C
5
coa 148 1 bwolAlpine
coah
coali
G
del
A
C/T
T
C/T
T
C
C/T
C/T
5
coal4817FtDavis
coah
coah
G
T
A
C
T
C/T
T
C
T
C/T
2
coal4818FtDavis
coah
coah
G
del
A
T
T
T
T
C
C
C
1
coal4819FtDavis
coah
coah
G
del
G
T
T
T
T
C
C
C
1
coal4820FtDavis 689G/T
coah
coah
A/G
del
A/G
C/T
T
T
T
C
C/T
C
6
coal 482 IFtDavis
coah
coah
G
del
A/G
T
T
T
T
C
C
C
2
coa 1 4822sofMarfa
coah
coah
A/G
T
A
C
T
T
T
C
T
C
1
coa 1 4823sofMarfa
coah
coah
G
T
A
C
T
T
T
C
C/T
c
1
coa 1 4824solMarfa
coah
coah
G
del
A/G
T
T
T
T
C
C
c
2
coa 1 4825sofMarfa
coah
AxC
A/G
T
A
C
T
T
A/T
C
T
c
2
coa 1 4826solMarfa
coah
coah
A/G
del
na
na
na
na
T
C
C/T
c
3
coa6829krn85. nLaZarca. rose
coah
coali
A/G
T
A
C
T
T
T
C
T
c
1
coa6830km83, nLaZarca, rose
coah
coah
A
T
A
C
T
T
T
C
T
c
0
coa6831krn85. nLaZarca, rose '
coah
coiih
A/G
T
A
c
T
T
T
C
T
c
2
coa 1 024 1 k 1 8 nDgo blue-pink "
BF
coah
G
T
A
c
T
T
T
C/T
T
c
2
coa 10242k 18 nDgo blue-pink ^
BF
coah
G
T
A
c
T
T
T
C/T
T
c
2
coal0503LaParr red, sweet Fr
BF
coah
G
T
A
c
T
C/T
T
C
T
C/T
2
coa l0504LaParr red,s\veet
coah
coah
A
T
A
c
T
T
T
C
T
C
0
coal0505LaPaiT violet Fr
BF
coah
G
T
A
c
T
T
T
C
T
c
0
coa 1 05 12LaParr red- orange Fr
BF
coali
G
T
A
c
T
T
T
C
T
C
0
coal0454LaParr. rose Fr
coah
hyb?
G
del
A
T
T
T
T
C
C/T
C
2
coal0455LaParr bluish Fr ^
BF
LaPar
G
del
A/G
T
T
T
T
C
C
C
4
coa 1 0456LaParr rose-purple
coah
hyb?
G
del
A
T
T
T
T
C
C/T
C
3
coal0459Guan rose-red.ro bio
coah
LaPar
G
del
A
T
T
T
T
C
C
c
1
coa 105 1 ILaParr red-orange Fr
coah
LaPar
G
del
A
T
T
T
T
C
C
C
1
coaBF14902LaParr blueFr
BF
matt
G
del
A
G
T
T
T
C
T
c
1
coaBF14903Tepeh blueFr
BF
mart
G
del
A
G
T
T
T
C
C/T
C
2
coaBF14904SPapa blueFr
BF
mart
G
del
A
G
T
T
T
C
T
C
1
mart5950 J. martinezii
mart
mart
G
T
A
G
T
T
T
C
T
C
0
mart5950 J. martinezii
mart
mart
G
T
A
G
T
T
T
C
T
C
0
‘ 240 AG; -603A/G; '503C/T; ‘^731 AG; -‘’308A/G, 665C/T; with ca. 20% C at site 196
74
Phytologia (Jan 19, 2017) 99(1)
The taxa of Dictyomorpha (Chytridiomycota, in praesens tempus)
Will H. Blackwell, Peter M. Letcher and Martha J. Powell
Biological Sciences, The University of Alabama, Tuscaloosa, AL 35487
ABSTRACT
Dictyomorpha (initially known, among Chytridiomycetes, as PringsheimieUa), an endoparasite of
types of ‘water molds’ (e.g. Achlya), is relatively unusual in being a heterothallic chytrid. As traditionally
recognized, Dictyomoipha belongs to Family Olpidiaceae, Order Chytridiales. The genus was long
considered monotypic, D. clioica the only taxon known. However, an additional variety (D. dioica var.
pytf liens is) was eventually described, seemingly based exclusively on occurrence in a different host
(Pythium). Without explanation, this variety w^as subsequently elevated (different author) to species. We
reviewed the mo, putative taxa of Dictyomorpfui in an attempt to detemiine whether varietal or specific
status is preferable. Based on apparent moqDhological distinctions evident in existing literature and
illustrations, tlie rank of species is supported, viz. Dictyomoipha dioica and D. pythiensis. We also
consider whether Dictyomorpha should remain in Phylum Chytridiomycota, or, rather, if this genus is
perhaps more appropriately placed in Phylum Cryptomycota (“Superphylum” Opisthosporidia).
Published on-line www, phytologia.org Phytologia 99(1): 74-82 (Jan 19, 2017). ISSN 030319430.
KEY WORDS: Achlya, aquatic fungi, chytrid, Dictyomorpha^ endoparasite, Nucleophaga, Olpidiaceae,
Oomycetes, Plasmophagus, Pringsheimiella, Pythium, resting spores, Rozella, sporangia, zoospores.
Dictyomorpha (originally Pringsheimiella) — an endobiotic, single-celled genus producing small,
posteriorly uniflagellate zoospores — has been considered a member of the Olpidiaceae; this family
contains holocaipic fonns (simple thallus converting, asexually, entirely to a sporangium), lacking
rhizoids (i.e., lacking “vegetative” staictures). Althougli tlie Olpidiaceae has traditionally been placed in
Order Chytridiales (Class Chytridiomycetes), recent placements of some members have indicated other
relationships (as will be discussed). The name Dictyomoipha (‘net-fomi”) would seem [incorrectly] to
imply a ‘network’ or ‘multicellularity;’ mutual compression of zoosporangia in [sorus-like] clusters in the
host (cf Mullins, 1961; Karling, 1977) — ^resultant from multiple zoospore infections (cf Couch, 1939;
Karling, 1977) — imparts this illusion. Dictyomoipha should not be confused witli Dictyiichus (name =
‘net-holder’ ), an unrelated genus [of Oomycetes] in which a single sporangium may contain a network of
(its own) zoospore-cysts (cf Blackwell and Powell, 1999). Karling (1977) — and previously Couch
(1939), ref. Pringsheimiella — noted that, superficially, Dictyomorpha may resemble [perhaps be mistaken
for] Dictyiichus (until one realizes that the “D/crywr/u/x-like” appearance of Dictyomoipha is the result of
a combination of the morphology of Dictyomoipha and its host, e.g., Achlya — and not simply the
consequence of morphological development of a single organism ).
Dictyomorpha (for many years thouglit to contain only D. dioica) was loiowTi as a parasite of
Achlya (A. ""flagellataf cf Couch, 1939; Mullins, 1961); D. dioica is relatively distinct among
Chytridiomycota in being heterothallic (apparently existing as moiphologically similar, male and female
strains). A new variety (D. dioica var. pythiensis) was later discovered in a species of Pythium (Sarkar
and Dayal, 1988). Dictyomorpha dioica was thouglit to be morphologically uniform, in spite of
recognition of tins additional variety (see Sarkar & Dayak 1988), how^ever, this variety was eventually
recognized as a species by Dick (2001) who provided no supporting evidence for his elevation of
taxonomic level. The zoospores of Dictyomorpha — and its resting spores (tliese fonned as the result of
sexual reproduction, by motile gametes seemingly identical to zoospores ) — ^bear resemblance to those of
Rozella, cf Mullins (1961). Rozella had been considered a genus of Chytridiomycota, but some species
Phytologia (Jan 19, 2017) 99(1)
75
are now classified elsewhere (discussed hereui) — ^raising questions as to correct phylum placement of
Dictyomorpha. Our study questions the unifomiity of Dictyonwrpha, examines potential taxa in the genus
(their ‘rank’), and reconsiders relationships of this genus among Fungi and related organisms.
TAXONOMIC HISTORY OF DICTYOMORPHA (Figures 1 - 20)
Dictyomorpha was described as a genus of Chytridiales (Mullins, 1961). Illustration (Fig. 1) of
[what turned out to be] sporangia of this organism [in its host, Achlya} is, however, traceable to
Pringsheim (1860, specimens from Genu any). Pringsheim, though, provided no legitmiate name for this
organism, incorrectly interpreting the motile cells he observed (his plate 22, fig. 5 and plate 23, fig. 3) as a
stage (antherozoids) in the life-cycle of Achlya [unrelated gejuis of Oomycetes]. Achlya and other
Saprolegniaceae do not possess flagellated gametes. Motile cells [actually zoospores] figured by
Pringsheim are uniflagellate (flagellum at or toward one pole), Figs, 1,3. Zoospores of Achlya and other
Saprolegniaceae are biflagellate {Achlya is laterally biflagellate). Motile cells do not seem to have been
illustrated by Cornu ( 1 872); however, the organism seen by him (similar to that illustrated by Pringsheim)
was placed in Cornu’s new genus, Woronimi (non-chytridiomycetous organism — classified in the
Plasmodiophoromycetes, e.g., Alexopoulos, 1962). Sparrow’s (1943, fig. 44A) illustration of the
organism seen by Cornu (1872) matches generally with that illustrated by Pringsheim (1860). Couch
(1939) — ^noted in Sparrow (1943) — described ‘Pringsheim’s organism,’ not as a Plasmodiophoromycete,
but more correctly as a chytridialean genus — under his proposed name, Pringsheimiella (acknowledging
Pringsheim’s illustration). Couch (1939), based on collections in North and South Carolina, accurately
described zoospores of Pringsheimiella as posteriorly uniflagellate. Couch, realizing that Pringsheimiella
had been known just in its asexTial phase, determined P, dioica (then the only taxon) to be heterothallic —
among the first members of the Chytridiales shown to be so — ^male and female strains necessary' for
sexual reproduction (and production of resting spores. Figs. 1 1-12). Couch noted potential physiological
(not morphological) differences between certain strains. Sparrow (1960) recognized Pringsheimiella
Couch (1939). Mullins (1961) was uncertain that the organisms seen by Pring.sheim (1860) and Couch
(1939) were the same; however, Karling’s (1977) illustration of this organism compares well with those
of Couch and Pringsheim. There is little doubt that Pringsheim’s fig. 1, plate 23, is of sporangia (in
Achlya) of what would be described as Pringsheimiella (Couch, 1939) and Dictyomorpha (Mullins,
1961 ), Mullins was concerned that Pringsheim didn’t observe [the zoospore as having] the lipoid body of
chytrid zoospores; however, certain of Pringsheim’s illustrations (plate 23, fig. 3) suggest this feature.
Mullins (1961) reviewed the taxonomy/nomenclature of Pringsheimiella Couch (1939),
concluding the generic name was preoccupied; Mullins indicated that ''Pringsheimielld' was employed by
Hohnel, in "‘1919” in vol. “17” of Ami. MycoL, as the name of an alga. Nielsen and Pedersen (1977)
noted that HohneFs use of this algal name was actually in 1920 (vol. 18). Regardless, because of
Hohnel’s prior usage, Pringsheimiella Couch (1939) is a later homonym (illegitimate). Mullins (1961)
supplied a legitimate, substitute name Dictyomorpha [nomen novnm] for Pringsheimiella Couch. Mullins
re-collected Dictyomorpha (Highlands, NC area) and restudied the life cycle — ^providing additional
description and illustrations (including zoospore variation, see Fig. 9), and depositing slide material
(additional to that of Couch, re; Pringsheimiella) in the UNC herbaiium. Still, only one species was
recognized in Dictyomorpha', this species, named D. 'dlioicd' by Mullins (1961), would seem to have
been transferred from Pringsheimiella (P. ^'dioica'd Couch, 1939). One might assume tliis species name
would be cited '^Dictyomoipha dioica (Couch) Mullins” — and it is so cited by Karling (1977) and Dick
(2001). However, Index Fimgonim currently (correctly we believe) lists the citation as ''Dictyomorpha
dioica Couch ex Mullins” — doubtless because Couch (1939) provided no Latin diagnosis when he
described genus Pringsheimiella and species P. dioica (relegating Couch to having 'proposed’ the epithet
""dioicd^ rather than legitimately publishing it). Mullins (1961) provided a combined, Latin genus/species
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Phytologia (Jan 19, 2017) 99(1)
description for Dictyomorpha/D. dioica, validating both. Authorship of the name D. dioica could, in fact,
be cited as either “Couch ex Mullins” or “Mullins” (cf International Code..., Article 46.5).
Pursuant to Mullins (1961), Dictomorpha was still thought to contain only D. dioica, without sub-
specific taxa, until Sarkar and Dayal (1988)» based on material from India, described D. dioica var.
pythiensis (see our Figs. 15-20) — occurring in Pythium aphanidermatum — automatically creating
Dictyomorpha dioica var. dioica [not mentioned by Sarkar and Dayal]. While attempts by Sarkar and
Dayal to hi feet hosts (including Adilya) other than Pythium aphanidemialum witli D. dioica var.
pythiensis were unsuccessfiil — var. pythiensis being apparently host-specific — ^they indicated a “close
morphological similarity” between their variety and typical D, dioica, noting no consistent morphological
differences; they felt, therefore, that var. pythiensis could not be justified as a new species. Since host
specificity was nonetheless considered important in deciphermg entities within the Olpidiaceae (Sparrow,
1960; Mullins and Barksdale, 1965), Sarkar and Dayal (1988) deemed varietal recognition appropriate.
As noted by Sarkar and Dayal, Mullins and Bai'ksdale ( 1965) demonstrated an increased host range for
Dictyomorpha dioica [i.e., var. dioica] , successful infections included a total of eight identified (and two
unidentified) species of Achlya (primarily in Subgenus Aclilya) — including the original host (A.
flagellata) — and also, Thraustotheca clavata\ Pythiiim was not included in their investigation.
Questionable evidence fi'om early literature (Pringsheim, 1860) suggested that D. dioica may have
occurred in Saprolegnia (cf. Mullins, 1961); however, Saprolegnia isolates tested (Mullins and
Barksdale, 1965) were immune to such infection. The ^^Saprolegnkf identified by Prmgsheim (1860, his
plate 22) was apparently a mixture of Achlya and Dictyuchus (the latter not involving Dictyomorpha).
In a nomenclatural summary, Dick (2001) — ^placing Dictyomorpha in Family Rozellopsidaceae,
Order R ozell ops i dales (Order "dnsertae sedis') — recognized two species, "^Dictyomorpha dioica (J. N.
Couch) J. T. Mullins” and ""Dictyornoipha pythiensis (N. Sarkai’ 8l R. Dayal) M, W. Dick, stat. nov.”
Proper author citation of D. dioica [i.e.. Couch ex Mullins] has already been discussed. Of concern is
Dick's (2001) recognition of var. pythiensis (Sarkar & Dayal, 1988) at species level, since Dick offered
no justification for this status change (no distinguishing features of the taxa were noted). As we
mentioned, Sarkar and Dayal had recognized ""pythiensis"" as a variety of D. dioica (not a separate species)
because ""pythiensis"' was based, by them, on host specificity — occurring in Pythium, not Achlya — rather
than on morphology. There was thus a need to determine if there are in faet morphologieal differenees
between the two alleged taxa within Dictyomorpha.
AT WHICH RANK SHOULD THE TAXA OF DICTYOMORPHA BE RECOGNIZED?
The question henee remains: Should the two ‘entities’ (var. dioica and var. pythiensis) within
Dictyomorpha dioica be considered varieties (Sarkar and Dayal, 1988) or species (Dick, 2001)? Dick
presented no evidence for his decision to recognize Dictomorpha dioica and D. pythiensis as distinet
species. If there is no reliable difference between these ‘taxa' other than host occupied (implied by Sarkar
and Dayal, 1988), varietal status would be (at most) the appropriate taxonomic category. Even if this ‘host
difference’ is accompanied by only one, minor, morphological difference, varietal status is perhaps still
preferable. But if there is separation of taxa by host infeeted andhy several morphological differences,
species recognition should be considered. Reexamination of literature (including illustrations) was
essential to this determination, since living material is not currently available; future collection of
Dictyomorpha is obviously important to further understanding of the genus.
Comparison of illustrations of [what eventually came to be known as] Dictyomorpha dioica in
Pringsheun (1860), Couch (1939), Mullins (1961), Karling (1977) and Sai'kar and Dayal (1988) —
reference our Figs. 1-20 — suggests (in addition to occurrence in mutually exclusive hosts) that
morphological differences do exist between “var. dioica"" and "'var. pythiensis.""' Eight (8) potential
differences we noted in these illustrations — ^not always congruent with statements in te.xt of the articles —
Phytologia (Jan 19, 2017) 99(1)
77
include: 1 . Shape of zoosporangium — “dioicaj' typically spherical (Figs. 1-2), except as altered by
mutual compression; ''pythiensisd' generally oval (Figs. 17-18). 2. Sporangial discharge "‘tube” —
''dioicar merely a papilla (Figs. 6-7); “pythiensls," occundng as an actual (sometimes somewhat
elongated) tube (Figs. 16-18). 3 . Number of sporangia in host cell — “dioicci2' often numerous (Fig. 1);
''pythiemis'^ ranging from one to eight, illustrated (Sarkar and DayaL 1988) as six or fewer (Figs. 17-18).
4 . Location of sporangia in host — ''dioicaj" occurring in vegetative (often distal/apical) portions of host-
hyphae (Figs. 1,2,6); ^'pythiensisl' occurring at various points in vegetative hyphae and, notably, in
oogonia (Figs. 17-18). 5 . Sporangial wall — ''dioica“ relatively tliin and pliable (Fig. 7); '‘pythiensis,”
firmer and more definite in shape (Figs. 17-18). 6. Zoospores — ''dioica,'"' illustrated (cf Fig. 3) as
typically somewhat elongated (Pringsheim, 1860) or irregular (spherical to elongate, e. g., Mullins, 1961;
Karling, 1977), illustrated (cf Fig. 15) as essentially spherical (Sarkar and Dayal, 1988,
though stated by them to be elongate). 7. Resting-spore outer wall — ‘JlioiccC roughened, undulate,
reticulate, or obscurely spiny (Figs. 11-14); ''pythiensisr more distinctly spiny (Fig. 20), although the
spines are typically small. We note that Raiding (1977) illustrated (see, for example, fig. 36 of his plate 8)
the outer resting-spore wall of ‘WpicaF D. dioica as more obviously (though still minutely ) spiny than did
eitlier Couch (1939) or Mullins (1961). 8. 'Extra' structure (‘ compartment' ) surrounding the already
double- walled resting spore(s)? — ''dioica,''' one to several resting spores contained (often loosely) in a
sometimes thick- walled, polygonal to square or rounded, extra ‘celF or ‘compartmenf (Figs. 11,12,14)
produced by the host (illustrated: Couch, 1939; Mullins, 1961; Karling, 1977); "^pythiensis," no extra
‘host compartmenf surrounds resting spores, though host-hyphae may form septa (Fig. 20) in response to
infection (cf Sarkar and Dayal, 1988).
Certainly, not all characters are distinguishable between ‘‘‘'dioica' and '"pythiensis." For example:
Zoosporangial, and resting-spore, diameters of ""dioica" were indicated (respectively) to be 15 to 20 pm,
and 15 to 17 pm (Mullins, 1961); for pythiensis^ tliese same parameters were (respectively) observ^ed at
12 to 20 pm, and 14.95 to 18.95 pm (Sarkar and Dayal, 1988). Regardless of precise form, the small
zoospores of the two taxa ai‘e also of similar dimensions (ca. 3 pm; cf Couch. 1939; Mullins, 1960;
Sarkar and Dayal, 1988). The resting spores (other than degree of ‘spiny' appearance of the outer wall)
are not only similar between the two taxa of Dictyomorpha, but reminiscent as well of the resting spores
of Rozella (to which Dictyomorpha may be related; cf Mullins, 1961, p. 386, last paragraph).
Characters (whether potentially distinguishing or not) perceived through study of literature are
subject to frirther investigation should live material of Dictyomorpha become available. Regardless,
sufficient morphological differences seem evident in various illustrations — in consort with delimitation
by host infected — ^to support recognition of the varieties of Dictyomoipha dioica — D. dioica var. dioica
and D. dioica var, pythiensis (Sarkar and Dayal, 1988) — as separate species (Dick, 2001, although Dick
gave no explanation for this change in taxonomic status). We tlms accept (duly noting here proper
authorship) two species within Dictyomorpha'. D. dioica J. N. Couch ex J. T. Mullins (1961) and D.
pythiensis (N. Sarkar & R. Dayal) M. W. Dick (2001). We camiot, though, concur with Dick's inclusion
of Dictyomorpha in the Rozellopsidaceae (Rozellopsidales); this category contains biflagellatc forms,
e.g., Rozellopsis. Zoospores of Dictyomorpha are definitely uniflagellate (Couch, 1939; Mullins, 1961;
Karling, 1977; Sarkar and Dayal, 1988), cf Figs. 3,8,9,15.
POSSIBLE SYSTEMATIC RELATIONSHIPS OF GENUS DICTYOMORPHA
Dictyomorpha — ^traditionally placed in the order Chytridiales [class Chytridiomycetes, phylum
Chytridiomycota] — was considered a member of the family Olpidiaceae (simple, holocarpic forms
lacking rhizoids). The Olpidiaceae included such seemingly similar genera as: Olpidium, Olpidiomorpha,
Rozella, Plasmophagus, Nucleophaga and Sphaerita (cf Sparrow, 1960; Karling, 1977). But molecular
infonnation has shed new light upon relationships of some Olpidiaceae. For example, certain species of
Olpidium place within the clade of Zygomycetes (James et al., 2006); and, species of both Rozella
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Phytologia (Jan 19, 2017) 99(1)
(Karpov et al., 2014) and Nucleophaga (Corsaro et al., 2014) have relationships within phylum
Ciy ptomycota (supeiphylum Opisthosporidia). Bearing, as it does, morphological similarity (of zoospores
and resting spores) to Rozella (cf, Mullins, 1961; Karling, 1977), Dictyomorpha could conceivably place
in the Ciyptomycota, rather than the Chytridiomycota; just how closely Dictyomorpha is related to
Rozella, remains to be determined. We do note that in Dictyomorjylia, in contrast to Rozella, the
sporangial walls are readily distinguishable from the wall of the host (cf Mullins, 1961, p, 386).
However, only molecular/genetic analysis will answer ultimate questions of generic and phylum
relationships. The puzzle of the systematic relationship of Dictyomorpha is, in fact, quite similar to that of
Plasmophagits (Blackwell et al., 2016). Unfortunately, these obligately parasitic organisms are not
generally available in culture collections — ^nor may they typically be cultured in the absence of their hosts
(cf Mullins, 1961; Mullins and Barksdale, 1965, re; Dictyomorpha dioica) — ^I'endering molecular
analysis elusive. Future collecting of such organisms — so that molecular analyses will have at least the
possibility of being performed — is essential to ultimate resolution of systematic problems. There is
continuing need for broad suiweys of ‘liydromy coflora” — such as that of Czeczuga (1995) in north-east
Poland — to “enrich our knowledge of biology of many aquatic fungi species.” Dictyomorpha dioica was
indeed found by Czeczuga, in one of 31 lalces sampled (the host for this organism, however, was not
indicated).
ACKNOWLEDGEMENTS
We thank tlie reviewers of this manuscript: Dr. Sonali Roychoudlniry, Patent Agent and Scientific
Consultant, New York; and Dr. Robert W. Roberson, Associate Professor, School of Life Sciences,
Arizona State University, We express appreciation to the following journals for permission to use or
reproduce illustrations; American Journal of Botany, Proceedings of the National Academy of Science,
India', and Journal of the Elisha Mitchell Scientific Society (currently Journal of the North Carolina
Academy of Science). We gratefully acknowledge NSF grant # 145561 1 for support of this research.
LITERATURE CITED
Alexopoulos, C. J. 1962. Introductory Mycology (2"^^^ ed.). Wiley; New York, London and Sydney.
Blackwell, W. H., P. M. Letcher and M. J. Powell. 2016. Reconsideration of the inclusiveness of genus
Plasmophagus (Chytridiomycota, posteris traditiis) based on morphology. Phytologia 98: 128-136.
Blackvs^ell, W. H. and M. J. Powell. 1999. Taxonomic summar}^ and reconsideration of the generic
concept of Dictyuch us. Mycotaxon 73: 247-256.
Cornu, M. 1872. Monographic des Saprolegniees; etude physiologique et systematique. Ann. Sci. Nat.
Bot, Ser. 5, 15: 1-198, pis. 1-7.
Corsaro, D., ,1. Walochnik, D. Venditti, K. D. Muller, B. Hauroder and R. Michel. 2014. Rediscovery of
Nucleophaga amoebae, a novel member ofthe Rozellomycota. Paristol. Res. 1 13: 4491-4498.
Couch, J. N. 1939. Heterothallism in the Chytridiales. J. Elisha Mitchell Soc. 55: 409-414, pi. 49.
Czeczuga, B. 1995. Hydromycoflora of thirty -one lakes in Elk Lake District and adjacent waters with
reference to the chemistry of the environment. Acta Mycol. 30: 49-63.
Dick, M. W. 2001. Sframinipilous Fungi. Khiwer Academic; Dordrecht, Boston and London.
Hohnel, F. von. 1920. Mykologische fragmente. Annales Mycologici 18: 71-98.
Index Fungomm (cuiTently updated online database of frmgal names), “www.indexfringorum.org”
Intemational Code of Nomenclature for algae, fungi, and plants. 2012. lAPT, Melbourne Code,
“www'.iapt-taxon.org/nomen/main.php”
James, T. Y., P. M. Letcher, J. E. Longcore, S. E. Mozley-Standridge, D. Porter, M. J. Powell, G. W.
Griffith and R. Vilgalys. 2006. A molecular phylogeny of the flagellated fungi (Chytridiomycota)
and description of a new Phylum (Blastocladiomycota). Mycologia 98: 860-871.
Karlmg, J. S. 1977. Chytridiomycetarum Iconographia. J. Cramer; Vaduz, Liechtenstein; and Lubrecht &
Cramer; Monti cello. New York.
Phytologia (Jan 19, 2017) 99(1)
19
Karpov, S. A., M. A. Mamkaeva, V. V. Aleoshin. E. Nassonova, O. Lilje and F. H. Gleason. 2014.
Moiphology, phylogeny, and ecology of the aphelids (Aplielidea, Opisthokonta) and proposal for
the new superphylum Opisthosporidia. Front. Microbiol. 5: 1 12. doi: 10.3389/fmicb.20 14.001 12.
Mullins, J. T. 1961. The life cycle and development of Dictyomorpha gen. nov. (formerly
Pringsheimielld), a genus of the aquatic fungi. Amer. J. Bot. 48: 377-387.
Mullins, J. T. and A. W. Barksdale. 1965. Parasitism of the chytiid Dictyomorpha dioica. Mycologia 57:
352-359.
Nielsen, R. and P. M. Pedersen. 1977. Separation of Syncoiyne reinkei nov. gen., nov. sp. from
Pringsheimiella sciitata (Chlorophyceae, Chaetophoraceae). Phycologia 16: 41 1-416.
Prmgsheim, N. 1860. Beitrage zur Morphologic und Systematic der Algen. IV. Nachtrage zur
Morphologic der Saprolegnieen. Jahrb. Wiss. Bot. 2: 205-236, pis. 22-25.
Sarkar, N. and R. Dayal. 1988. A new variety of Dictyomorpha dioica (Couch) Mullins. Proc. Nat. Acad.
Sci. India 58 (Sec. B, III): 403-406.
Sparrow, F. K. 1943. Aquatic Phy corny cetes. Exclusive of the Saprolegniaceae and Pythium. Univ.
Michigan Press, Ann Arbor; Elumphrey Milford, London; and Oxford Univ. Press.
Sparrow, F. K. 1960. Aquatic Phy corny cetes, 2"^ revised edition. Univ. Michigan Press, Ann Arbor.
80
Phytologia (Jan 19, 2017) 99(1)
Figs. 1-8: Dictyomorpha dioica. 1: Sporangia (Sp), generally spherieal in form, in host (Achlya);
zoospores released at tip of host filament (arrow). 2: Diseharged sporangia (arrow). 3: Variable
(often elongate) shape of posteriorly uniflagellate zoospores. 4: Zoospores infeeting host
{Achlya) by their apieal ends. 5: Young thalli (Th) developing in host. 6: Maturing, and also
empty, sporangia inside apieal portion of host hypha. 7: Maturing sporangia (Sp); note exit-
papilla (arrow). 8: Mature sporangium; zoospores released, through papilla, laterally from host
filament. Figs. 1-3 after Pringsheim (1860), 4-5 after Mullins (1961), 6 after Coueh (1939) and
Mullins (1961), 7-8 after Mullins (1961).
Phytologia (Jan 19, 2017) 99(1)
81
Figs. 9-14: Dictyomorpha dioica. 9: Range of zoospore form. 10: ‘Zoospores’ fusing, as
gametes, to form zygote. 11-14: Resting spores (RS) in various maturation stages (in hyphae of
host, Achlya); note ‘extra eells’ (‘host eompartments’ = He), eaeh surrounding one to several
resting spores (Figs. 11, 12, 14); wall of ‘host eompartments’ sometimes thiekened (12); outer
resting-spore wall roughened, retieulate or ‘undulate’ (11-12), sometimes sub-spiny (13). Figs. 9-
10 after Mullins (1961), 11-12 after Coueh (1939), 13 after Mullins (1961), 14 based generally
on Coueh (1939) and Karling (1977), among others.
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Phytologia (Jan 19, 2017) 99(1)
Figs. 15-20: Dictyomorpha pythiensis. 15: Zoospore. 16: Sporangium (Sp) in host (Pythium),
exit-tube forming (arrow). 17-18: Emptied sporangia (Sp) in host oogonium (Og); sporangia
generally oval, exit-tubes persistent. 19: Resting spores (RS) in host oogonia. 20: Resting spores
(RS), inside host hypha, exhibiting minutely but distinetly spiny walls; speeial ‘host
eompartments’ (potentially enclosing resting spores) lacking, but extra hyphal septa may form
(arrow). Figs. 15-20 after Sarkar and Dayal (1988).
Phytologia (Jan 19, 2017) 99(1)
83
Mandevilla torosa (Apocynaceae), treated as having two allopatric intergrading varieties in Mexico
Billie L. Turner
Plant Resources Center The University of Texas Austin, TX 78712
billie.turner@austin.utexas.edu
ABSTRACT
Mandevilla coulteri S. Wats, is treated as a variety within the widespread M. torosa, the former
largely confined to north-central Mexico, but passing into var. torosa southwards.
KEY WORDS: Apocynaceae, Mandevilla, MexicoPublished on-line www.phytologia.org Phytologia
99(1): 83-85 (Jan 19, 2017). ISSN 030319430.
Mandevilla torosa (Jacq.) Woodson, the Type from Jamaica, with populations extending into
southern Mexico, is treated as composed of two intergrading varieties, a more southern typical var. torosa
and a more northern var. coulteri, the latter largely confined to Coahuila, Nuevo Leon and Tamaulipas
but grading into var. torosa south- wards; this dichotomy was first proposed by Williams (1999) in his
doctoral thesis but not published. Unfortunately he applied the varietal name “karwinskii” to the more
northern elements, the latter typified by a Karwinski collection from southern Mexico (probably Oaxaca).
He should have adopted the varietal name “coulteri,” for the northern populations, which is typified by a
Coulter collection from the state of Coahuila.
I have more or less adopted the key to the two varieties provided by Williams, but with the
addition of leaf shapes:
Mandevilla torosa (Jacq.) Woodson, Ann. Missouri Bot. Card. 19: 64 1932.
Key to varieties
1. Corolla tubes mostly 4-6 mm long; leaves mostly obovate, or rounded
at their apices; plants typically vine-like var. torosa
1. Corolla tubes mostly 7-9 mm long; leaves mostly elliptic with acute apices;
plants perennial herbs or subshrubs var. coulteri
var. coulteri (S. Wats) B.L. Turner, var. nov.
Based upon Echites coulteri S. Wats., Proc. Amer. Acad. Arts 18: 113. 1883.
The name is typified by Coulter 957, this collected in the state of Coahuila, S. of Saltillo,
according to Williams (1999). There are some 60 specimens of the variety at LL- TEX, all remarkably
alike and possessing the characters attributed to var. coulteri by the present author. Williams applied the
name var. karwinskii to all of these sheets, largely because he had not examined the type concerned;
Alvorado-Cadenas and Morales (2014) correctly note its synonymy under their concept of Mandevilla
torosa; they also placed var. coulteri in synonymy under M. torosa, which belies the taxonomy proposed
herein.
Distribution of the two taxa in Mexico, along with intermediates, is show in Fig. 1. W illi ams
mapped, but did not annotate or name the intermediate sheets. Those sheets which I have accepted as
intermediates (and mapped accordingly) follow:
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Phytologia (Jan 19, 2017) 99(1)
TAMAULIPAS (two collections): Hidalgo, Hinton et al. 24709', 11 mi W of Victoria. Graham &
Johnston 4133.
SAN LUIS POTOSI: Barkley et al. 854; Irving 167 [both collections near Cd. de Maiz].
QUERETARO: (7 sheets, all intermediate) Carranza 930', Carranza & Silva 5873a', Fernandez &
Rzedowski 3425', Rubio 1866, 1250', Sendn 1042; Zannidio & Carranza 6651.
Alvorado-Cadenas and Morales (2014) noted two collections of M. torosa from Veracruz that I
have not examined. I have mapped these as var. torosa, but these too might be intermediates. Indeed,
with DNA analysis it is possible that typical Mandevilla torosa (in Mexico) will be found confined to the
Yucatan Peninsula and that intermediates between these and vai\ coulteri are deserving of formal
recognition.
It should be noted that Morales (1998) stated ''Mandevilla karninskii is closely related to
M, torosa but can be recognized by its ver\^ narrowly elliptic (or almost linear) to spatulate leaf blades and
mucroulate to rarely acute leaf apices, its usually sub-erect habit, and the usually continuous to obscurely
moniliform follicles.” He did not clearly delineate the two taxon, either morphologically or
geographically, as perceived by Williams, or the present author.
LITERATURE CITED
Alvarado-Cardenas, L.O. and Morales, J.F. 2014. El gcmro Mandevilla (Apocynaceae: Apocynoideae,
Mesechiteae) en Mexico. Botanical Sciences 92: 59-79.
Morales. J.F. 1998. A synopsis of the genus Mandevilla (Apoeynaceae) in Mexico and Central America.
Brittonia 50: 214-232.
Williams, J.K. 1999. A phylogenetic and taxonomic study of the Apocynaceae, subfamily Apocynoideae
of Mexico, with a synopsis of subfamily Plumerioideae. Doctoral Diss„ Univ. of Texas, Austin, 546
pp.
ACKNOWLEDGEMENTS
Thanks to LL-TEX for specimens examined and to Jana Kos for editorial input.
Phytologia (Jan 19, 2017) 99(1)
85
Fig. 1. Distribution of Mandevilla torosa in Mexico
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