Jan 2001 * ip' i v fl irS ■w f *»jp. I ara 1 jlt flmilVi • rJvr« *3 if- * Oh & 65 Years Lower Higher Lower 15(60%) 10(40%) Religiosity Higher 14(29%) 34(71%) Total 29(40%) 44(60%) Chi Square = 6.53 ( 1 df); P = 011 Kendall’s tau - b = .30; p = .01 1 6 Mullins and Brackett Table 2c. Crosstabular Analysis: Marital Adjustment x Religiosity x Sex Marital Adjustment Male Lower Higher Lower 29 (60%) 19 (40%) Religiosity Higher 24(39%) 38(61%) Total 53(48%) 57(52%) Chi Square = 5.1 1 (ldf); p = .034 Kendall’s tau - b = .21; p = .021 Female Lower Higher Lower 26(62%) 16(38%) Religiosity Higher 31(48%) 34(52%) Total 57(53%) 50(47%) Chi Square = 2.07 (ldf); p = .150 Kendall’s tau - b =. 137; p = .145 7 Religiosity and Marital Adjustment Table 2d. Crosstabular Analysis: Marital Adjustment x Religiosity x Race Marital Adjustment Blacks Lower Higher Lower 10(53%) 9 (47%) Religiosity Higher 23(61%) 15(39%) Total 33(58%) 24(42%) Chi Square = 0.32 (ldf); p = .584 Kendall’s tau - b = .-.08; p = .571 Whites Lower Higher Lower 42(63%) 25(37%) Religiosity Higher 30(37%) 52663%) Total 72(48%) 77(52%) Chi Square =10.07 (ldf); p = -002 Kendall’s tau - b =.260; p = .001 8 Mullins and Brackett The association, however, between religiosity and marital adjustment for those aged 65 and older is strong and statistically significant (jf = 6.53, ldf, p< 02), while the association between these two variables for those under age 65 is relatively weaker (x2=1.35, ldf, p=.3 1). Similarly, among the men in the study, there is a strong positive and statistically significant association between religiosity and marital adjustment = 5.11, ldf, p<.024). Among the women the association is in the same positive direction, but weaker and nonsignificant (x‘ = 2.07, ldf, p=. 15) (Table 2c). The racial differences are striking. Among Whites (x2 = 10.07, ldf, p< 002), marital adjustment is both statistically significant and strong (Table 2d). This is not the case among the Blacks (x2 =0.32, ldf, p=58); religiosity essentially is unrelated to expressed marital adjustment among the African American in this study. DISCUSSION The results here point to a relatively weak, but clear, link between religious commitment, involvement, ideology and related aspects, and a greater degree of what here has been called marital adjustment, especially among older white males. Booth et.al (1995) stated that “social scientists have a long history of research attempting to explain how religious sentiment direct social action” (p. 661). They go on to say that “more recently. . . scholars have questioned religion’s capacity to serve as a socially integrative force in contemporary society” (p. 661). The results suggest there may be a cohort, racial and sex difference in the way that religion influences marriage. Durkheim (1951) in an extension of his early sociological examination on suicide emphasized that both religion and marriage are independent integrative forces in the reduction of destructive tendencies, or conversely in the enhancement of constructive tendencies (though this was not tested). Religion as a factor in social integration could influence marital quality on a structural level in that it provides a context within which the marriage or the relationship is defined. The religious context could be the frame on which the marriage as an institution rests. On a social psychological level the religious involvement, used as means for the interpretation of the meaning of life or as a model for behavior, could serve to guide the individual in his or her relationship with the spouse. Seminal work in the past several decades (D’Antonio, Newman & Wright, 1982; Thomas, 1988a, 1988b; Thomas & Cornwall, 1990; Thornton, 1985) suggests on a theoretical level that marriage and religion work as both social control and social support mechanisms. Thomas and Cornwall (1990) state: “The social support dimension emphasizes that religion supports family life through norms that encourage love, family solidarity, and marital satisfaction [while] the social control mechanism emphasizes the impact of religion as constraining behavior” (p. 986). Could it be that the lack of impact of religion on marital adjustment among the younger subgroup in the present study indicates the suspected impact of what many believe is a more secular society? Results for the older subgroup show the weak, but significant, impact of religion on the marital relationship. This may be part of the explanation, but there are other ideas to consider. It could be that those who are older have found that religion is a social institution that contains provisions for a structured life, 9 Religiosity and Marital Adjustment or religion is a mechanism that provides meaning to life within the context of the relationships one has. Religion is now a more relevant aspect of life. It is well documented that church or synagogue membership increases with age (U.S. Bureau of the Census, 1993). Those who are younger simply may not find the time for, or do not see the current relevance of, religion in the lives they lead. The race differences in the religiosity/marital adjustment relationship are no less interesting. The fact that there was a significant relationship between religiosity and marital adjustment among Whites, but not Blacks, is especially intriguing. The African American church, historically, has been especially important in the maintenance of what is a conservative and traditional set of values that cross-cuts African-American life. Mindel, Habenstein and Wright (1998) point out that “churches have been strong, powerful, and responsible sources of family support” (p. 376) that provides religious support and solace, as well as of employment and financial assistance. The results here may point to a potential erosion of the impact of the church and religion in general in the substance of the marital relationship among African-Americans. Among Whites, church and religion has been more varied in its impact, due at least in part to both the greater denominational diversity and the intermingling of religion with national identity, e.g., Anglo-Saxon Protestant, Irish Catholic, Greek Orthodox. Within ethnic/national categories there is at least some religious variability. The whites in the current study, however, are predominantly Protestant — Southern Baptist at that — with an essentially fundamentalist orientation. Protestant fundamentalism, according to Hunter (1983, 1985, 1987), has five broad aspects that distinguishes it from other denominations: literal interpretation of the scriptures, non-acceptance of religious pluralism, pursuit of a personal experience with God, opposition to “secular humanism,” and endorsement of conservative political goals. There is currently much emphasis within the more conservative wing of Protestantism on the importance of church, qua fundamentalist-interpreted religious orientation, on the stability of marriage, and the harmony and satisfaction within the marriage. The results here, i.e., the religiosity-marital adjustment relationship, may be reflecting either the real, or the expected, degree of marital harmony as a result of the high degree of stated religiosity within the fundamentalist tradition. On a social psychological level another issue is of possible relevance, also. Is communication, and therefore the marriage, influenced by the relevance of religion and religious activities? An interesting study by Snow and Compton suggests that “when religion is important to a spouse, it may affect marital communication by increasing empathy for one’s partner and decreasing hostile communication” (p. 985). Echoing some things stated in the introduction of this paper, the development of a shared reality about many aspects of life, including religion, is crucial to a positive marital relationship. The results of this study also point toward a gender difference. While males and females both demonstrated a positive relationship between religiosity and marital adjustment, this was much stronger for males. The presumed role of men in marriage, head of household, is supported through religious teachings, particularly the fundamentalist teachings followed by the majority of the respondents. In following the traditional roles in the home, men receive encouragement and support from religion. In this way the cultural models are reinforced with religion fulfilling a social support function. Men thus receive congruent messages about their sex appropriate roles in the marriage. The continuing prominence of groups such as Promise Keepers and initiatives such as that by the Southern Baptist Convention indicating women’s 10 Mullins and Brackett subordinate status to men encourage the traditional model of marital adjustment. This is particularly true with regard to decision making and roles in the home. This is a model with which men are likely to be more comfortable than are women. Religion then reinforces their adjustment. For women, religion provides messages of both social support and social control. Women’s traditional marital role of nurturer and helper is consistent with religious doctrines, particularly those of fundamentalist protestants. Religion, however, is not women’s only source of information about marital roles; secular models are readily available. The ideals of equality espoused by feminism and inculcated into modern society are inconsistent with fundamentalist perspectives. In this situation women may view religion as a model that provides social support for traditional roles and exercises social control over alternate roles. Thus women’s marital roles have been subject to more alternative ideas and competing sources of information. Additionally, the weaker relationship between religiosity and marital adjustment among women might relate to social psychological elements, such as how much shared reality there is between the couple. While men may take their relationships for granted, women tend to engage in more monitoring. Women’s tendency to analyze their relationships might make them more aware of any areas of difficulty, particularly potential communication differences. Burgess, et al (1963) suggest that successful communication and subsequent agreement on issues is a hallmark of marital adjustment. Scholars repeatedly suggest that males and females use and interpret language differently so that conversations are not experienced the same by both partners (Tanner, 1990 and 1994; Lakoff, 1975). If males and females experience their marital communication differently it is reasonable that their levels of adjustment differ or are influenced by different social and relationship variables. Berger (1967) proposes that the links between religion and family life remain strong because they are both part of the private sphere, having influence there rather than in the public sphere. The most profound changes in recent years in women’s roles have come in their contributions in the public sphere, i.e. their employment behaviors. Despite changes in their employment status, women continue to perform more traditional tasks in the home. As Heaton and Cornwall (1989) reported family behavior is much more influenced by religion than is economic behavior. Thus in the private sphere traditional models of marriage remain strong. Since religion and family are key institutions in the private sphere, it is not unexpected that religiosity and marital adjustment are linked. Given their greater burden in home tasks, it is not surprising that the relationship between religiosity and marital adjustment is weaker among women. The link between religiosity and marital adjustment in this study was not unexpected. Montgomery is a community in which religious affiliation is important and encouraged. Religion can be a touchstone for couples, providing guidance and shared values upon which to construct a lifetime relationship. For many partners the same religious orientation is a prerequisite for marriage. It should be noted that this study’s population was intact married couples. The fact that they are still together indicates these are more conventional couples. They may hold more traditional values, particularly with regard to working through issues to preserve the relationship. The most interesting elements of the present study warrant further investigation. By including additional variables, it would be possible to understand other factors related to Religiosity and Marital Adjustment marital adjustment, as well as the strength of religiosity relative to other factors. These alternate explanations should be explored with regard to younger couples and African Americans. Further study may uncover whether the gender difference in the strength of the relationship between religiosity and marital adjustment is best explained by gender role orientation, relationship dynamics, or other factors. Finally, exploring this topic in a population of fewer fundamentalist believers may result in different findings and should be pursued. LITERATURE CITED Bellah, R., Madsen, R., Sullivan, S., Swindler, A., & Tipton, S. (1985). Habits of the heart: Individualism and commitment in American life. New York: Harper and Row^ Berger, P. 1967. The Sacred Canopy. Garden City, NY: Anchor Books. Booth, A. Johnson, D., Branamann, A., & Sica, A. (1995) Belief and behavior: Does religion matter in today’s marriage? Journal of Marriage and the Family, 57, 661-673. Burgess, E.W., Locke, H.J., & Thomas, M.M. (1963). The family: From institution to companionship (3rd ed.). New York: American Book. Busby, D., Christenson, C., Crane, D., & Larson, J. (1995). A revision of the dyadic adjustment scale for use with distressed and nondistressed couples: Construct hierarchy and multidimensional scales. Journal of Marriage and Family Therapy. 21, 298-308. Chi, S.K., & Houseknecht, S.K. (1985). Protestant fundamentalism and marital success: A comparative approach. Sociology and Social Research, 69, 351-373. D’Antonio, W.V., Newman, W.M., & Wright, S.A. (1982). Religion and family life: How social scientists view the relationship. Journal for the Scientific Study of Religion, 21, 218-225. Durkheim, E. (1951). Suicide: A study in sociology. New York: Free Press. Etheridge, F. M. (1979). Varieties of fundamentalism: A conceptual and empirical analysis of two Protestant denominations. The Sociological Quarterly, 20, 49-48. Faulkner, J., & DeJong, G. (1966). Religiosity in 5-D: An empiricial analysis. Social Forces, 45, 246-254. Glock, C. (1962). On the study of religious commitment. Religious Education, 57, 98-1 10. Heaton, T., & Cornwall, M. (1989). Religious group variation in the socioeconomic status and family behavior of women. Journal for the Scientific Study of Religion, 28, 283- 299. Heaton, T. & Pratt, E. L. (1990). The effects of religions homogamy on marital satisfaction and stability. Journal of Family Issues, 1 1, 191-207. Hunter, J.D. (1983). American evangelics. New Brunswick, NJ.. Rutgers University Press. Hunter, J.D. (1985). Conservative Protestantism. In P.E. Hammond (ed.). The sacred in a secular society (pp. 1 50- 1 66). Berkeley: University of California Press. Hunter, J.D. (1987). Evangelicalism: The coming generation. Chicago: University of Chicago Press. Lakoff, R. (1975). Language and Woman’s Place. New York: Harper & Row. Lauer, R. (1995). Social problems. Madison, WI: Brown & Benchmark. 12 Mullins and Brackett Lewis, R. A., & Spanier, G.B. (1979). Theorizing about the quality and stability of marriage. In W. H. Bum, R. Hill, F.I. Nye, & I. L. Reiss (eds.). Contemporary theories about the family (Vol. 1) (pp. 268-294). New York: Free Press. Mindel, C.H., Habenstein, R.W., Wright, Jr., (1998). Ethnic families in America: Pattern and variations (3rd ed.). New York: Elsevier Shehan, C. L., & Bock, E. W. (1990). Religious heterogamy, religiosity, and marital happiness: The case of Catholics. Journal of Marriage and the Family. 52. 73-78. Sillars. A.L., & Scott, M. D. (1983). Interpersonal perception between intimates. Human Communication Research, 10, 153-176. Snow, T.S., & Compton, W.C. (1996). Marital satisfaction and communication in fundamentalist protestant marriages. Psychological Reports, 78, 979-985. Spanier, G. B. (1976). Measuring dyadic adjustment: New scales for assessing the quality of marriage and other dyads. Journal of Marriage and the Family, 38, 15-23. Tannen, D. (1990). You Just Don’t Understand: Women and Men in conversation. New York: William Morrow, Ballantine. Tannen, D. (1994). Talking From 9 to 5. New York: William Morrow & Co., Inc. Thomas, D. L. (1988a). Future prospects for religion and family studies: The Mormon case. In D. L. Thomas (ed). The religion and family connection: Social science perspectives (pp. 357-382) Provo, UT: Religious Studies Center, Brigham Young University. Thomas, D.L. (1988b). The religion and family connection: Social science perspectives. Provo, UT: Religious Studies Center, Brigham Young University. Thomas, D. L. & Cornwall, M. (1990). Religion and family in the 1 980’s: Discovery and development. Journal of Marriage and the Family. 52, 983-992. Thornton, A. (1985). Reciprocal influences of family and religion in a changing world. Journal of Marriage and the Family. 47, 381-397 United States Bureau of the Census (1993). Statistical Abstract of the United States (113 ed.). Washington, DC: U.S. Government Printing Office. *Please address correspondence to Larry C. Mullins, Ph.D., School of Liberal Arts, P.O. Box 244023, Auburn University Montgomery, Montgomery, AL 36124-4023. 13 Journal of the Alabama Academy of Science, Vol. 72, No. 1, January 2001. INTERVASCULAR PIT STRUCTURE IN SELECTED SPECIES OF THYMELAEACEAE Roland R. Dute1, Michael E. Miller1,2 Robert R. Carollo1 'Department of Biological Sciences, 2AU Research Instrumentation Facility Auburn University Auburn, AL 36849 ABSTRACT Intervascular pit pairs in woods of Gnidia cajfra , Dirca palustris, and two species of Pimelea were observed with both light and scanning electron microscopy. The resulting images were compared with pits from Daphne spp. of a previous study. Unlike Daphne , the pit membranes of the other genera lack tori. Pit apertures of Gnidia and Pimelea were slit-like, whereas those of Daphne and Dirca tended toward circular. Vestures were present in intervascular pit pairs Gnidia , Pimelea , and, to a lesser extent, in Dirca, but absent from the Daphne species investigated in this study. Hypotheses regarding the function of tori and vestures are discussed. INTRODUCTION Pit pairs located between water-conducting cells (tracheary elements) in wood provide a low-resistance pathway for water movement but at the same time must inhibit gas embolisms from passing cell to cell. There are two basic mechanisms to accomplish these functions. In many conifers and Ginkgo, the pit membrane separating the pits consists of microfibrils associated with large openings. In the center of this screen-like barrier is an impermeable circle of wall material, the torus (Thomas, 1969). The openings through the pit membrane serve for water movement. In the presence of embolisms, however, the pit membrane is deflected or aspirated such that the torus blocks the aperture into the neighboring cell thus impeding movement of bubbles (Zimmermann, 1983). In contrast, pit membranes in most dicotyledonous woods consist of microfibrils associated with very small (or no) visible openings and no torus. Since the pressure differential needed to force an air bubble through a screen increases with decreasing size of the openings, the intact pit membrane effectively blocks movement of most embolisms (Zimmermann, 1983). There are, however, a very few species of dicotyledons whose pit membranes have both the dicot arrangement of microfibrils as well as a torus. Among them are species of the genus Daphne (a shrub common to Eurasia), and it is this genus which has been the subject of intensive investigation in this laboratory. The presence of tori in membranes of Daphne was first noted by Ohtani and Ishida 14 Dute, et al. (1978) Dute et al. (1990) detailed torus development in this genus and hypothesized that such a structure prevented membrane rupture during aspiration. Both Ohtani (1983) and Dute et al. (1992) noted that not all species of Daphne possessed a torus, but in fact presence or absence of this feature was determined by systematics. Specifically, species of section Mezereum lacked a torus (Dute et al., 1996). This, same study noted tori in wood of two of three subgenera of Wikstroemia, a sister genus to Daphne. At the same time, tori were absent from wood of Drapetes and Edgeworthia, genera within the same subfamily (Thymelaeoideae) as Daphne. Table 1 presents an abbreviated classification of the family Thymelaeaceae according to Engler (1964). Those genera within the subfamily Thymelaeoideae which have been investigated are indicated with asterisks. Table 1. An abbreviated classification of the Thymelaeaceae adapted from Engler (1964). The genera marked with an asterisk have been previously studied in this laboratory. The genera listed in boldface are examined in the present study. Family - Thymelaeaceae Subfamily Tribe Genus Gonystyloideae Gonystylus, Amyxa, Aetoxylon Aquilarioideae Microsemmataeae Microsemma Solmsieae Solmsia, Deltaria Octolepideae Octolepis Aquilarieae Aquilaria, Gryinops Gilgiodaphnoideae Gilgiodaphne Thymelaeoideae Dicranolepideae Linostoma, Lophostoma, Dicranolepsis, Craterosiphon, Synaptolepsis Phalerieae Phaleria, Peddiea Daphneae *Wikstroemia, Dendrostellera, Daphnopsis, Dirca, *Daphne, * Edgeworthia Thymelaeeae Thymelaea, Gnidia, Struthiola, Passerina, Kelleria, * Drapetes, Pimelea 15 Intervascular Pit Structure Observations of these genera indicate that the torus-bearing pit membrane and the apertures work as a functional unit. When a torus is present, the aperture is small in diameter (smaller than the torus) and is circular to slightly elliptical in outline. In some species lacking a torus, pit apertures may also be circular, whereas in other species, the apertures are slit-like (Dute et al., 1996). The association of circular apertures with torus-bearing pit membranes has been noted by other authors (Beck et al., 1982; see also discussion in Morrow and Dute, 1998). Vestures, wall protuberances associated with pits, are listed as a feature of the Thymelaeaceae (Bailey, 1933; Jansen et al., 1998; Metcalfe & Chalk, 1950). These structures have been hypothesized to reduce membrane rupture during aspiration (Zweypfenning, 1978) or to prevent formation of (gas) embolisms or aid in their resorption (Carlquist, 1988). Thus a knowledge of vesture structure and systematic occurrence as well as an understanding of the torus could be important to comprehending safety features in wood. This manuscript represents a continuation of work on intervascular pits of the Thymelaeaceae. Specifically, it provides a characterization of pits in Gnidia caffra (Meisn.) Gilg, Dirca palustris L., and two species of Pimelea with emphases on the presence or absence of a torus, the outline of the pit apertures, and presence or absence of vestures. Images of pits from Daphne gnidioides, D. collina, and D. retusa are included for comparative purposes. MATERIALS AND METHODS The species used in this study are listed in Table 2. All specimens represent air-dried herbarium samples and with one exception were obtained from the Auburn University Herbarium (AUA). Daphne gnidioides samples included in this study were used in a previous publication (Dute et al., 1992). Only Daphne gnidioides was re-investigated in detail although individual micrographs of other Daphne species are presented. Branch segments of 3—6 mm diameter were split longitudinally to expose either radial or tangential surfaces. Samples were then affixed to aluminum stubs with double stick carbon tape and sputter-coated with gold- palladium. Observations were made with a Zeiss Digital Scanning Microscope (DSM 940) at voltages of 5, 10, and 15 kV. Measurements of SEM material were made using the system included with the microscope. All measurements were determined by counts of 25 unless otherwise noted. SEM observations were supplemented with light microscopy. Herbarium material was treated and embedded in Spurr's resin (Spurr, 1969) according to the procedure of Dute et al. (1990). Thin sections of 1.5—3 pm were cut on an ultramicrotome and heat-fixed to glass slides. The sections were stained in one of three ways: 1) 0.5% toluidine blue O (TBO), 2) 2% aqueous KMnC>4, or 3) KMnC>4 followed by TBO. The latter staining procedure provided higher contrast than either of the other two methods. Apparently, KMn04, in its role of oxidizing agent, exposed more potential binding sites for the subsequently applied TBO molecules. Once stained, specimens destined for light microscopy were covered by mounting medium and a coverslip. Photographs were taken using T-Max 100 film. 16 Dute, et al. Table 2. Sources of wood specimens examined in this study. Taxon Herbarium or City Date of Collection Collector(s) No. Pimelea arenaria ( 1 ) AUA 1 1 Oct 1979 Cooper & Nickerson 61 13 - (2) AUA 22 Dec 1979 Wilkinson & Nickerson 6386 Pimelea prostrata AUA 26 Jan 1980 Nickerson & Nickerson 6458 Dirca palustris (1) AUA 27 Mar 1969 Krai 34029 - (2) AUA 27 Mar 1969 Krai 34034 - (3) AUA 19 Mar 1989 Diamond & Freeman 573 1 Gnidia cajfra ( 1 ) AUA Jan 2000 Boyd & Davis s.n. - (2) AUA Jan 2000 Boyd & Davis s.n. Daphne gnidiodes K 26 Jul 1960 Khan, Prance & Ratcliffe 255 RESULTS General Information Fig. 1 is an overall, radial longitudinal view of a water-conducting cell or vessel member in the wood of Dirca palustris. The large opening at the top, known as a perforation, is actually located on the inclined end wall of the cell. A similar hole is located at the bottom of the cell, but is slanted in the opposite direction and is therefore not exposed to the observer. Vessel members are stacked end to end with their end walls overlapping and their perforations juxtaposed. This series of cells is referred to as a vessel. Tire top and bottom elements of the vessel have only a single lower and upper perforation, respectively. Thus a vessel acts as a closed system. In addition, a second type of water-conducting cell, the tracheid, can also be present. This cell type differs from vessel members by the absence of perforations, but in other 17 Intervascular Pit Structure respects (i.e. pitting) is similar. Daphne, Dirca, and Pimelea have tracheids as well as vessel members; Gnidia has only the latter. For convenience, the general term “tracheary element” (Esau, 1965) will be used when referring to both kinds of conducting elements unless a distinction is warranted. Lateral walls of the tracheary elements are pitted (Fig. 1), and it is the pits between tracheary elements (the so-called bordered pit pairs, Fig. 2) which provide a pathway for lateral movement of water. Fig. 1 shows these pits in face view for D. palustris. A detailed view of a similar aspect is provided in Fig. 3 for Gnidia caffra. Pit pairs can be viewed at different levels. For, example, letter A in Fig. 3 indicates the outer surface of a pit border. The hole or aperture through that border opens into the lumen or cavity of the tracheary element (which, for the cell in question, is below the surface of this figure and thus not visible). Letter B represents a pit membrane overlying the border. Letter C identifies the inner or lumen surface of the other pit border of the pair (with its own aperture). Thus the pit membrane is inserted between two pit borders. Fig. 4 provides a cross-sectional view of a pit pair of Daphne collina (at right angles to Fig. 3). The letters indicate the same surfaces as in the previous figure. In this instance the pit membrane is displaced (aspirated) to one side and covers the outer opening of an aperture. Membrane displacement is a typical occurrence in air-dried wood. Pit Membrane Structure Figs. 5, 6, 7 & 8 present pit membrane structure in Daphne species for comparative purposes. In the center of the pit membrane is a thickening known as a torus. This thickening is found on both sides of the pit membrane and can be seen in cross section even with the light microscope (Fig. 7). In face view the torus is circular with a diameter greater than that of the associated pit apertures (Table 3). Detailed views of such pit membranes indicate the torus to be an impermeable structure, whereas the remainder of the pit membrane (the margo), which surrounds the torus, is fibrillar (Figs. 6 & 8). Although present, fibrils are not always observed in air-dried pit membranes as wound material released by surrounding cells as they die impregnates the pit membranes and obscures the fibrils (Dute et al., 1992; Morrow & Dute, 1999; Schmitt & Liese, 1990). Detailed views of intervascular pit membranes in D. palustris (Figs. 9 & 10), both species of Pimelea (Figs. 1 1, 12), and G. caffra (Fig. 13) show no evidence of a torus. Pit Aperture Structure Species of Gnidia and Pimelea used in this study have irregularities known as vestures along the rims of their pit apertures (Figs. 1 1 & 13). Daphne gnidiodes does not (Fig. 5), and for the most part, Dirca palustris does not, although sometimes very small amounts of vesturing (best described as obscure) can be observed (Fig. 14). Vesture morphology of the Pimelea spp. differs from that of G. caffra. In the former, the vestures appear as irregularities at the very edge of the aperture (Fig. 1 1), whereas in Gnidia the apertures are peglike and cover part of the outer (non-lumen) surface of the pit cavity as well as the aperture rim (Fig. 13). Interestingly, fiber pits of Dirca, Gnidia, and Pimelea species observed in this study (as well those of Daphne gnidioides ) all have well-developed vestures. The ratio of the short axis to the long axis of intervascular pit apertures was used as a measure of aperture circularity. Table 3 presents results for all genera investigated in this study. From the data, it is evident that apertures of D. palustris and D. gnidioides more closely 18 Dute, et al. approach a circular shape than those apertures of P. prostata, P. arenaria, and G. caffra. Table 3. Dimensions of pit membranes, pit apertures, and tori. All measurements are in micrometers and are taken from the non-lumen side of the pit border. Means are based on 25 measurements. species pit diameter long axis of aperture short axis of aperture circularity ratio torus diameter Pimelea prostata 5.24 (4.37- 6.07) 1.89(1.27- 2.61) 0.56 (0.42- 0.81) 0.30 none P. arenaria ( 1 ) 5.46 (4.23— 6.77) 2.43 (1.76— 3.60) 0.71 (0.5- 1.20) 0.29 none P. arenaria (2) 5.14(4.23- 6.00) 3.02 (2.04— 4.23) 1.10(0.84- 1.41) 0.36 none Dirca palustris (1) 5.57 (4.58- 7.05) 1.60(1.20- 2.04) 1.16(0.63- 1.69) 0.73 none D. palustris (2) 6.68(5.71- 7.58) 1.62 (1.05- 2.22) 1.27 (0.88— 1.58) 0.78 none D. palustris (3) 6.31 (5.29- 8.00) 2.19(1.12- 3.64) 1.39(0.91- 1.88) 0.63 none Daphne gnidiodes 6.58(5.41- 8.82) 1.62 (1.05- 2.22) 1.15 (0.88- 1.58) 0.71 2.77 (2.28- 3.29) Gnidia caffra d) 3.95 (2.96- 5.29) 2.22(1.83- 3.45) 0.58 (0.40- 0.84) 0.26 none G. caffra (2) 4.88 (4.23- 5.71) 2.33 (1.54— 2.96) 0.65 (0.45- 1.12) 0.28 none 19 Intervascular Pit Structure Key to labelling: AP = pit aperture; M = margo; P = pit; PE = perforation; T = torus; V = vesture. Figure 1. Radial longitudinal section (SEM) of Dirca palustris wood showing a vessel member with a perforation and pits. Scale bar =20 pm. Figure 2. Tracheary elements (A) of D. palustris pulled away from neighboring elements (B) to show bordered pit connections. Scale bar = 50 pm. 20 Dute, et al. Figure 3. Detailed view of Gnidia caffra showing pit pairs at different levels. "A" is the outer surface of a pit border, "B", the pit membrane; "C", the lumen surface of the pit border. Scale bar = 5 pm. Figure 4. Sectional view of pit pair (from Daphne collina) normal to the surface of Fig. 3. Letters A, B, and C represent the same surfaces as in the previous Fig. Scale bar = 2 pm. Figure 5. Face view of pits in Daphne gnidioides. Note the torus and its larger diameter compared to the nearby pit aperture. Scale bar = 2 pm. Figure 6. Oblique view of torus-bearing membrane of Daphne retusa. The torus and margo regions are easily distinguished. Scale bar = 1 pm. Figure 7. Light micrograph of sectional view through pits of Daphne gnidioides. Scale bar = 5 pm. Figure 8. Detail of torus-bearing membrane (of Daphne alpina) in face view. Note the fibrillar nature of the margo. Scale = 1 pm. 21 Intervascular Pit Structure Figure 9. Figure 10. Figure 1 1 . Figure 12. Figure 13. Figure 14. Pit membranes of Dirca palustris. No torus is present. Dark circle in center of membranes represents subtending aperture. Scale bar = 2 pm. Light micrograph of Dirca palustris wood showing pit pairs with pit membranes in sectional aspect. Scale bar = 5 pm. Pimelea prostrata pit membranes (right) and pit borders (left). Note vestures on apertures and tearing of pit membranes (unlabeled arrow). Scale bar - 2 pm. Sectional view of P. arenaria pit membranes. Scale bar = 5 pm. Pit vestures and portion of a pit membrane (asterisk), Gnidia caffra. Scale bar = 2 pm. Pit apertures of Dirca palustris with obscure vesturing (arrow). Scale bar = 5 pm. n Dute, et al. DISCUSSION Torus— distribution and function Torus distribution in the Thymelaeaceae is of some value to systematics. In a study of 22 spp. of Daphne, Dute el al. (1992) observed 19 spp. with tori. This structure was absent from three spp. of the section Mezereum. A later study of the sister genus Wiskstroemia (Dute et al., 1996) showed tori to be present in the subgenera Diplomorpha and Daphnimorpha , but absent from the subgenus Wikstroemia. The presence of tori in the Thymelaeaceae must be tightly circumscribed. Although Daphne and Wikstroemia are within the same tribe ( Daphneae , Table 1), so are Edgeworthia and Dirca but neither of the latter two genera possess tori (Dute et al, 1996; the present study). The more distantly related genera, Drapetes , Gnidia, and Pimelea (different tribe, same subfamily) also have no tori. Torus function in Daphne pit membranes is indicated by its structure. Tori are typically circular with a diameter greater than that of the aperture (i.e. the greatest diameter of the aperture). In a developmental study of pit membranes of Daphne odora and D. cneorum , the tori were of greater diameter than the pit apertures and completely occluded them during aspiration (Dute et al., 1990). The torus and aperture diameters of D. gnidioides from this study (2.8 jam torus; 1.6 pm aperture) are not out of line with measurements of D. cneorum (3.6; 1.8) and D. odora (3.6; 1.4) from the previous investigation. It is hypothesized that the torus in dicots decreases possibility of rupture of the pit membrane during aspiration (Dute et al., 1990). D. odora pit membranes, dried after torus removal, were ruptured at the site where the torus once overlay the pit aperture. Similar ruptures were observed in air-dried material of D. mezereum that never had a torus. While pit membrane damage could well have occurred during processing for SEM or due to heat of the electron beam, the fact remains that aspirated pit membranes tear where they overlay the pit aperture, whereas membranes with tori do not. The weakness of non-torus bearing membranes can be observed in Pimelea membranes (Fig. 1 1) of the present study. It appears as if pit membranes with circular tori are associated with more or less circular apertures, but the converse need not be true. Beck et al. (1982), writing with regard to gymnosperm wood, observed that circular apertures associated with circular tori would be "more highly adaptive" than slit-like apertures and provide an effective seal. By contrast, a less specialized homogeneous membrane would probably be adaptive in pits with slit-like apertures. However, they then cite Wright (1928), who states, "the presence of a round pore does not necessarily entail the development of a torus." It appears as if the same can be said for angiosperms. Clearly, in this study the circularity ratio is much less for pit apertures of Pimelea and Gnidia than for Daphne and Dirca. The former genera have slit-like apertures and have no tori. Daphne with a more circular aperture has a torus, but Dirca whose aperture is of a similar circularity, does not. As mentioned earlier, Dirca is closely related to Daphne and perhaps is preadapted by virtue of its circular aperture to use a torus should one arise in the course of evolution. Further evidence for correlation of torus development and aperture outline comes from Daphne aurantiaca and D. genkwa (Dute et al., 1996). Both species have both narrow and wide tracheary elements. The narrow ones have circular apertures and pit membranes with well-developed tori, whereas the larger elements have elliptical to slit-like apertures associated with tori of variable development or with no torus at all. 23 Intervascular Pit Structure Vestures— function and systematic distribution Vestures, in the restricted sense, represent wall protuberances associated with pits (Jansen et al., 1998). Zweypfenning (1978) hypothesized that these structures reduced membrane deflection during aspiration and decreased the possibility of membrane tearing. But as Carlquist (1988) rightly indicates, vestures and warts (protuberances on the surfaces of lumen walls) are the same structures in different locations. Thus, any hypothesis put forward must explain both features. According to Carlquist (1982), warts and vestures, by increasing surface area, could increase bonding of water molecules at the cell surface (lumen surface). This in turn would prevent breaking of water columns and formation of vapor bubbles. At this point, we would disagree with both hypotheses. True, in our observations it appears as if vestures are associated with slit-like apertures and could in theory substitute for the torus as regards protection for the pit membrane. However, construction of the vestures in Pimelea (Fig. 1 1, this study) is not such as to reduce membrane deflection and certainly does not prevent membrane rupture. Also, vestured pit membranes are present in fibers of all species examined in this study. These are cells which conduct little, if any, water. Ohtani (1987), noting the evidence for vestures (warts) in septate fibers and parenchyma cells, stated that vestures have no function associated specifically with pits of conductive elements, but are simply "formed by an oversupply of wall material from the protoplast at the final stage of secondary wall formation process." Whatever their function, vestures are considered to be a feature of the wood of the Thymelaeaceae (Bailey 1933; Metcalfe & Chalk, 1950). As recent examples, vestures have been reported for vessel or fiber pits of some Daphne spp. ( Dute el al., 1992; Ohtani & Ishida, 1976), vessel pits of Dirca and Ovidia (Record & Hess, 1943), and Aquilaria agallecha (Rao & Dayal, 1992). We have confirmed the presence of vestures in vessel pits of Dirca , although their presence is inconsistent, and when present, obscure. To the best of our knowledge, vestures in Gnidia are reported for the first time. There is conflicting evidence for the presence of vestures in various species of Pimelea. Ohtani et al. (1983) indicated that warts (and by extension vestures) are absent from both vessels and fibers of P. aridula, P. gnidia, P. oreophila, and P. traversii, yet a year later Ohtani et al. (1984) located vestures in P. gnidia, P. oreophila, P. pseudo-lyalli, and P. traversii. The presence of vestures in Pimelea has been extended to the species P. prostata and P. arenaria by the present investigation. ACKNOWLEDGEMENTS We wish to thank Drs. Robert Boyd and Micheal Davis for procuring specimens of Guidia for the herbarium and Mr. Curtis Hansen, herbarium curator, for his assistance. LITERATURE CITED Bailey, I. W. 1933. The cambium and its derivative tissues: VIII. Structure, distribution, and diagnostic significance of vestured pits in dicotyledons. J. Arnold Arbor. 14: 259—273. Beck C. B., K. Coy, and R. Schmid. 1982. Observations on the fine structure of Callixylon wood. Amer. J. Bot. 69: 54—76. 24 Dute, et al. Carlquist, S. 1982. Wood anatomy of Onagraceae: further species; root anatomy; significance of vestured pits and allied structures in dicotyledons. Ann. Missouri Bot. Gard. 69: 755-769. Carlquist, S. 1988. Comparative Wood Anatomy. Springer-Verlag, Berlin. Dute, R.R., J.D. Freeman, F. Henning, and L.D. Barnard. 1996. Intervascular pit membrane structure in Daphne and Wikstroemia— Systematic implications. IAWA J. 17: 161 — 181. Dute, R.R., A.E. Rushing, and J.D. Freeman. 1992. Survey of intervessel pit membrane structure in Daphne species. IAWA Bull. 13: 113—123. Dute, R.R., A.E. Rushing, and J.W. Perry. 1990. Torus structure and development in species of Daphne. IAWABull.il: 401-412. Engler, A. 1964. Syllabus der Pflanzenfamilien. Gebriider Bomtraeger, Berlin. Esau, K. 1965. Plant Anatomy, 2nd edition. Wiley, New York. Jansen, S., E. Smets, and P. Baas. 1998. Vestures in woody plants: a review. IAWA J. 19: 347- 382. Metcalfe, C.R. and L. Chalk. 1950. Anatomy of the Dicotyledons. Vol. 2. Oxford at the Clarendon Press. Morrow, A.C. and R.R. Dute. 1998. Development and structure of pit membranes in the rhizome of the woody fern Botrychium dissectum. IAWA J. 19: 429—441. Morrow, A. C. and R.R. Dute. 1999. Electron microscopic investigation of the coating found on torus-bearing pit membranes of Botrychium dissectum, the common grape fern. IAWA J. 20: 359-373. Ohtani, J. 1983. SEM investigation on the micromorphology of vessel wall sculptures. Research Bulletins of the College Experiment Forests. College of Agriculture, Hokkaido University 40: 323—386. Ohtani, J. 1987. Vestures in septate wood fibres. IAWA Bull. 8: 59—67. Ohtani, J. and S. Ishida. 1976. Study on the pit of wood cells using scanning electron microscopy. Report 5. Vestured pits in Japanese dicotyledonous woods. Research Bulletins of the College Experiment Forests, College of Agriculture, Hokkaido University. 33: 407—436. Ohtani, J. and S. Ishida. 1978. Pit membrane with torus in dicotyledonous woods. J. Jap. Wood Res. Soc. 24: 673-675. Ohtani, J., B.A. Meylan, and B.G. Butterfield. 1983. Occurrence of warts in the vessel elements and fibres of New Zealand woods. New Zealand J. Bot. 21: 359—372. Ohtani, J., B.A. Meylan, and B. G. Butterfield. 1984. A note on vestures on helical thickenings. IAWA Bull. 5: 9-11. Rao, K. R. and R. Dayal. 1992. Tire secondary xylem of Aquilaria agallocha (Thymelaeaceae) and the formation of ‘agar’. IAWA Bull. 13: 163—172. Record, S.J. and R.W. Hess. 1943. Timbers of the New World. Yale University Press, New Haven. Schmitt, U. and W. Liese 1990. Wound reaction of the parenchyma in Betula. IAWA Bull. 1 1 : 413-420. Spurr, A.R. 1969. A low viscosity epoxy resin embedding medium for electron microscopy. J. Ultrastruct. Res. 26: 31—45. 25 Intervascular Pit Structure Thomas, R.J. 1969. The ultrastructure of southern pine bordered pit membranes as revealed by specialized drying techniques. Wood and Fiber 1: 1 1 0 — 1 23 . Wright, J.G. 1928. The pit-closing membrane in the wood of the lower gymnosperms. Proc. Trans. R. Soc. Canada, Ser 3, 22: 63—95. Zimmermann, M.H. 1983. Xylem Structure and the Ascent of Sap. Spring-Verlag, Berlin. Zweypfenning, R.C.V.J. 1978. A hypothesis on the function of vestured pits. IAWA Bull. 1978/1: 13-15. 26 Journal of the Alabama Academy of Science, Vol. 72, No. 1, January 2001. THE DEVELOPMENTAL TOXICITY OF MECLIZINE USING THE FROG EMBRYO TERATOGENESIS ASSAY-XENOPUS (FETAX) Andrea Wolfe and James Rayburn Department of Biology Jacksonville State University 700 Pelham Rd. N Jacksonville, AL 36265-1602 ABSTRACT Meclizine, an antihistamine used to treat nausea and vomiting, was proven a potent teratogen for rats, causing abnormalities such as cleft palate, small mouth, short limbs, receded lower jaw, and uncalcified vertebral bodies in rat offspring (King, 1963). Frog Embryo Teratogenesis Assay-A \enopus (FETAX) has been proposed to serve as an indicator of potential human teratogenic hazards (Bantle et al., 1994). The purpose of this research was to determine how well FETAX could predict the known mammalian developmental toxicity of meclizine. The procedure involved the exposure of varying concentrations of meclizine to early life stages of Xenopus laevis, a South African clawed frog, for 96 hours. An average lethal concentration to cause 50% death in the test population (96 hour-LC50) was calculated as 4.62 mg/L. In addition to the 96 hour-LC50, an average concentration to induce 50% malformation (96 hour-EC50) was calculated as 0.974 mg/L. The average teratogenic index (TI) for meclizine was calculated as 4.74. According to ASTM standards (ASTM, 1991), a TI (96 hour-LC50/ 96 hour-EC50) of 1.5 or greater indicates teratogenic risk. Therefore, findings in FETAX with meclizine are consistent with the mammalian literature. INTRODUCTION Meclizine is a piperizine derivative antihistamine (Figure 1. Haraguchi et al., 1997) used commonly to treat nausea and vomiting related to motion sickness (Drug Info. American Hospital Formulary Service, 1998; Merck Index, 1996). It can also be used to treat vertigo associated with diseases of the vestibular system (Physicians Desk Reference, 1995). The mechanism of action for meclizine is not precisely known, but is “thought to be related to its central anticholinergic action. It is also believed that an action on the medullary chemoreceptive trigger zone may be involved in the antiemetic action of the drug” (Drug Information for the Health Care Provider, 1987). Meclizine has an oral LD50 of 1600 mg/kg and an intraperitoneal LD50 of 659 mg/kg (Sigma Chemical Company, Material Safety Data Sheet). Meclizine is a class B 27 Developmental Toxicity of Meclizine pregnancy drug. This means as long as benefits outweigh the potential risks, pregnant women may take the drug. Meclizine is generally considered a “safe” drug for pregnancy (Drug Info. American Hospital Formulary Service, 1998), but some epidemiological studies have suggested that meclizine could pose potential risk to the unborn fetus (Lione and Scialli, 1996). Meclizine Figure 1. Structure of meclizine Meclizine has been a “suspect” teratogen for years, as indicated by Schardein (1993). Some human studies in the United Kingdom showed that women treated with meclizine (plus vitamin B12) had offspring with defects, while other studies showed meclizine having no teratogenic effect (Schardein, 1993). In one case, a woman medicated with meclizine plus pyridoxine (vitamin B12) in the fourth week of gestation, had an infant with a unilateral limb defect (Jaffe et al., 1975). In another case cited by Walker (1974) a woman took meclizine and another antiemetic and had a child with iniencephaly and spina bifida. Bokesoy et al. (1983), cited a case in which a woman treated with meclizine throughout pregnancy, but primarily in the first trimester, had a child with severe micrognathia (abnormally small jaw), small mouth, widened nasal bridge, receded lower lip, cleft palate, and hypoplastic tongue and mandible. King (1963), analyzed meclizine-induced malformations such as cleft palate, micromelia (short limbs), microstomia (small mouth), brachygnathia (receded lower jaw), and uncalcified vertebral bodies in rat offspring. According to Schardein (1993), “animal species successfully demonstrate the potential for teratogenic effect for all known human teratogens.” With this in mind, meclizine could very well be considered a potential human teratogen. More studies need to be performed, however, to further demonstrate the teratogenicity of the drug meclizine. A conserved genetic program controls embryonic development, therefore in animal assays such as FETAX data can be extrapolated to other species (Bantle, 1995). Bantle (1995) indicated that FETAX was initially designed as an indicator of potential human' developmental hazards. This assay is well suited for complex mixtures and has been ranging from the effects of pond water from Minnesota and Vermont (Fort et al., 1999) to the benefits of ascorbic acid in preventing paraquat embryotoxicity (Vismara et al., 2001). 28 Wolfe and Rayburn The in vitro assay performed by Bantle et al. (1994), established that FETAX could be used to detect teratogens in mammals. This research was performed to determine if FETAX could correctly determine the developmental hazard of meclizine. This further validation of FETAX for the determination of developmental health hazards demonstrated its use in predicting the teratogenic hazards of chemicals. MATERIALS AND METHODS Frogs were obtained from Xenopus I (Dexter, MI). The drug meclizine, CAS Registry number 569-65-3, was obtained from Sigma Chemical Co. (St. Louis, MO). FETAX assay procedure followed the ASTM Standard Guide for the conduct of FETAX (ASTM, 1991). A reconstituted, mineralized water (FETAX solution) was used as the diluent, as described in the ASTM guide. Two replicates of 20 embryos each were placed into covered plastic Petri dishes for each test concentration. Four control plastic Petri dishes contained FETAX solution only. Embryos used were from the same clutch for each individual experiment. Three definitive experiments were run using meclizine with FETAX. Stock solutions of meclizine were made by dissolving 10 mg of meclizine in 1 L of FETAX solution by stirring and gentle heating. Afterwards, pH was adjusted (between 6-8) as necessary, then diluted into appropriate concentrations. Control and test solutions were then transferred to plastic Petri dishes and 20 frog embryos in small cell blastula stage were placed into plastic dishes with 8 mLs of test solution. Embryos were first examined under a dissecting microscope to be certain they were viable. Embryos were placed in an incubator at 24°C +. 1 . Dead embryos were removed and solutions were changed at 24, 48, and 72 hour intervals. After changing solutions, remaining embryos were counted. At the 96 hour interval surviving embryos were sedated with MS222, counted, and examined for defects then placed in formaldehyde solution, and later in isopropanol for length measurements. Length measurements of the embryos were made using Image Pro Analysis System. Values for 96 hour-LC50 (concentration required to kill 50%), 96 hour-EC50 (concentration where 50% of the embryos are malformed) and TI values (96 hour-LC50/96 hour-EC50) were also calculated using Litchfield-Wilcoxon probit analysis. Minimum Concentration to Inhibit Growth (MCIG) was determined using a grouped T-test. RESULTS Table 1 indicates the control results from the three experiments, which averaged 4.16 % mortality and 10.88% malformation. Mortality rate was below 5% for each of the three experiments. Malformation rate was more variable than mortality. However, only experiment 2 had a higher malformation rate than acceptable by the ASTM guide and all three experiments demonstrated similar results. 29 Developmental Toxicity of Meclizine Table 1. Meclizine Control Results Test # Mortality Malformation 1 5% 9.2% (4/80) (7/76) 2 3.75% 19.5% (3/80) (15/77) 3 3.75% 3.95% (3/80) (3/77) Average 4.16% 10.88% (10/240) (25/230) Table 2 contained the 96 hour-LC50, 96 hour-EC50, and TI of meclizine. The average 96 hour-LC50 and 96 hour-EC50 was 4.62 and 0.974 respectively, in mg/L. The average teratogenic index (TI) was 4.74. For experiment 1 no mortality above 50 % was achieved, so the 96 hour-LC50 and the TI was estimated not calculated. The averages for the 96 hour-LC50 and TI did not include experiment 1 . Table 2. Meclizine Test Results Test # 96 hour-LC50 (mg/L) 96 hour-EC50 (malformation) (mg/L) TI 1 >5 0.732 >6.8 0.13-4.2 2 6.02 1.18 5.1 0.86-41.98 0.044-31.62 3 3.21 1.01 3.2 0.36-28.33 0.16-6.30 Average 4.62* 0.974 4.74* ‘Does not include test 1 . ‘Does not include test 1 . 30 Wolfe and Rayburn The concentration-response curves represented in panel A of figures 2, 3, and 4 showed probit graphs of the test response for each of the three experiments. For Figure 2 panel A the mortality curve was different from the other two experiments because concentrations higher than 5 mg/L were not tested. Figures 3 and 4 panel A show extremely repeatable results from different clutches of embryos. Both mortality and malformation show a good linear probit concentration-response. The slopes for all malformation curves were approximately the same. Growth curves shown in panel B of figures 2, 3, and 4 indicated that growth is significantly inhibited between 90 and 155% of the 96 hour-LC50 (MCIG). At low concentrations there was apparently no effect on growth. Experiment 1 is the only experiment in which growth increased at low concentrations, however, this growth was not significant. The Minimum Concentration to Inhibit Growth (MCIG), shown in panel B, usually indicated the highest concentration, and was the only concentration significantly different from control. Figure 2. Panel A is the concentration-response mortality (O) and malformation (•) curves for meclizine from Experiment 1. Response is given in probit values— a probit of 5=50%. Panel B is embryo length measurements at 96 hours after exposure to meclizine— concentration is expressed as % of average LC50 vaiue length is expressed as % of mean control embryo length. * indicates significant difference on T-test (MCIG) p< 0.05. Figure 3. Panel A is the concentration-response mortality (O) and malformation (•) curves for meclizine from Experiment 2. Response is given in probit values— a probit of 5=50%. Panel B is embryo length measurements at 96 hours after exposure to meclizine— concentration is expressed as % of average LC50 value length is expressed as % of mean control embryo length. * indicates significant difference on T-test (MCIG) p< 0.05. Figure 4. Panel A is the concentration-response mortality (O) and malformation (•) curves for meclizine from Experiment 3. Response is given in probit values— a probit of 5=50%. Panel B is embryo length measurements at 96 hours after exposure to meclizine— concentration is expressed as % of average LC50 value length is expressed as % of mean control embryo length. * indicates significant difference on T-test (MCIG) p<0.05. 31 % Mean Control length Response (PROBIT) Developmental Toxicity of Meclizine Figure 2 Experiment 1 Probit Graph Experiment 1 Growth Graph 32 % Mean Control length Response (PROBIT) Wolfe and Rayburn Figure 3 Experiment 2 Probit Graph Experiment 2 Growth Graph 33 % Mean Control length Response (PROBIT) Developmental Toxicity of Meclizine Figure 4 Experiment 3 Probit Graph Experiment 3 Growth Graph 34 Wolfe and Rayburn Figure 5 contained embryos from the same test in three different concentrations. Embryo A was a control embryo with no malformation. Embryo B was from a 0.5 mg/L test concentration of meclizine, and embryo C was from a 5 mg/L test concentration. Figure 6 portrayed an embryo from a test concentration similar to that of embryo C in figure 5, these two embryos had similar malformations. The malformations seen such as gut and eye malformations, and multiple edema were remarkably similar in each experiment, indicating that meclizine causes specific types of abnormalities at increasing concentrations. Figure 5. Three embryos from meclizine test concentrations. A-Control, no malformations B-0.5 mg/L, edema, gut miscoiling, malformed tail, eye malformations C-5 mg/L, severe edema, malformed eyes, miscoiled gut, axial malformations 35 Developmental Toxicity of Meclizine Figure 6. An embryo from similar high concentration of a different test. The embryo has severe edema, malformed eyes, miscoded gut, and axial malformations. DISCUSSION The averages of all three experiments fell well within the acceptable ranges for the ASTM guide for the conduct of FETAX. The malformation percentage for experiment 2, however, was above acceptable limits. A possible cause for the discrepancy in the malformation averages could be explained by the subjective nature of the malformation scoring process. Each embryo was examined under a microscope and judging 36 Wolfe and Rayburn malformations was purely up to the person looking at the embryo. Experiment 2 may have been judged more severely than the other two experiments, or the embryos may have had a high malformation rate. The other two experiments had percentages well within the 10% range set forth by ASTM guidelines, showing that malformations in the meclizine test concentrations were not naturally occurring. The increased control mortality in experiment 2 did not appear to be a problem, as results from all three experiments were very similar as indicated by figures 2, 3, and 4. For experiment 1, fifty- percent mortality was not achieved, so a 96 hour-LC50 or TI could not be calculated for this experiment. An estimation was taken for the two values based upon the highest concentration tested. The 96 hour-LC50 was greater than 5 mg/L and the TI was estimated by taking the ratio of the estimated 96 hour-LC50 to the calculated 96 hour-EC50, giving a TI of greater than 6.8. In experiment 2, the 96 hour-LC50 was 6.02 mg/L, a number in agreement with the estimation from experiment 1. The 96hr-LC50 for test 3 was a little lower. More death occurred than expected in this experiment, however 95% confidence intervals were overlapping. The 96 hour-EC50 for all three experiments were within the same range, a good indication that the malformations noted are caused by meclizine, and not a one-time occurrence. Table 2 also shows that all numbers from the 96 hour-LC50 and 96 hour-EC50 fall within the ninety-five percent confidence limits for each of the three experiments. The average TI for the three experiments was 4.74, not far below the estimated TI for experiment 1. For all three experiments, the TI was above 1.5, showing that meclizine has potential to be a teratogen (ASTM, 1991). A TI of 4.74 indicated a moderate to high teratogenic risk for this drug. In figure 5 the first embryo, A, was a control embryo. It had no edema, normally set eyes, properly coiled gut, and a straight tail. Embryo B was from an intermediate concentration. It had some edema, eye malformations, and though not visible in the picture, it also had some gut miscoiling, and tail abnormality. The third embryo, C, from the highest test concentration, was severely malformed, with a great amount of edema, small eyes, miscoiled gut, and a curved axial alignment. The malformations of the embryo in figure 6 were very similar to the malformations of the third embryo (C) in figure 5. This property of repeatable results confirmed that meclizine was a teratogen in FETAX. CONCLUSION Meclizine causes severe malformations, gut and eye malformations, and multiple edema in FETAX, as evidenced by the embryos in figures 5 and 6. The data presented in this paper support literature that meclizine is a potential direct-acting human developmental toxicant. Lastly, these results are further validation that FETAX can serve as a good indicator of potential human teratogenic risk. ACKNOWLEDGEMENTS We thank Jacksonville State University Student-Faculty Research Grant for supporting this research. We also thank Randa Aladdin, Heather Howell, and Darryl Pendergrass for their technical assistance. 37 Developmental Toxicity of Meclizine LITERATURE CITED American Society for Testing Materials (ASTM). 1991. Standard Guide for Conducting the Frog Embryo Teratogeneisis Assccy -Xenopus (FETAX). E 1439-91. Philadelphia, PA. Bantle, J. 1995. FETAX-A Developmental Toxicity Assay Using Frog Embryos. In: Fundamentals of Aquatic Toxicology, Effects, Environmental Fate, and Risk Assessment. 2nd Edition. Ed. Gary Rand, Taylor and Francis. Washington D.C. Bantle, J., D. Burton, D. Dawson, J. Dumont, R. Finch, D. Fort, G. Linder, J. Rayburn, D. Buchwalter, A. Gaudet-Hull, M. Maurice, S. Turley. 1994. FETAX Interlaboratory Validation Study: Phase II Testing. Environmental Toxicology and Chemistry Vol. 13, No. 10:1629-1637. Bokesoy, I, Aksuyek, E. Deniz. 1983. Oromandibular Limb Hypogenesis/Hanhart’s Syndrome: Possible drug Influence on the Malformation. Clinical Genetics. 24(l):47-49. Drug Info. American Hospital Formulary Service. 1998. p. 2410-2411. Drug Information for the Health Care Provider. 17th ed. 1987. p. 1146-1147. Fort, D. T. Propst, E. Stover, J. Helgen, R. Levey, K. Gallagher, J. Burkhart. 1999. Effects of Pond Water, Sediment, and Sediment extracts from Minnesota and Vermont, USA, on Early Development and Metamorphosis of Xenopus. Environmental Toxicology and Chemistry. Vol. 18, issue 12:2305-2315. Haraguchi, K, K. Ito, H. Kotaki, Y. Sawada, T. Iga.1997. Prediction of Drug-Induced Catalepsy Based on Dopamine Dl, D2, and Muscarinic Acetylcholine Receptor Occupancies. Drug Metabolism and Disposition. 25:675-684. Jaffe, P, M. Liberman, I. McFadyen, H. Valman. 1975. Incidence of Congenital Limb- reduction Deformities. Lancet. 1:526-527. King, C. 1963. Teratogenic Effects of Meclizine Hydrochloride on the Rat. Science. 141: 353-355. Lione, A. and A. Scialli. 1996. The Developmental Toxicity of the HI Histamine Antagonists. Reprod. Toxicol. 10(4): 247-255. Merck Index. 1996. p. 984. Physicians Desk Reference-Generics. 1st ed. 1995. p. 1779-1780. Schardein, J. 1993. Chemically Induced Birth Defects. 2nd ed. Revised and expanded, pp. 71, 347, 446, 448-450. Vismara C., G. Vailati, and R. Bacchetta. 2001. Reduction in Paraquat Embryotoxicity by Ascorbic Acid in Xenopus laevis. Aquatic Toxicology. Vol. 51, issue 3:293-303. Walker, F. 1974. Familial Spina Bifida Associated with Antiemetic Ingestion in the First Semester. Birth Defects, 10:17-21. 38 Journal of the Alabama Academy of Science, Vol. 72, No. 1, January 2001. PTERIDOPHYTES OF NORTHEAST ALABAMA AND ADJACENT HIGHLANDS III: OPHIOGLOSSALES AND POLYPODIALES (Aspleniaceae to Dennstaedtiaceae) Daniel D. Spaulding Anniston Museum of Natural History Anniston, AL 36202 J. Mark Ballard Jordan, Jones, & Goulding, Inc. Tucker, GA 30084 R. David Whetstone Jacksonville State University Jacksonville, AL 36265 Illustrated by: Marion Montgomery Anniston Museum of Natural History Anniston, AL 36202 INTRODUCTION Members of Ophioglossales and Polypodiales belong to the division Polypodiophyta and are often called “true ferns.” The order Ophioglossales in Alabama include the adder’s-tongue ferns ( Ophioglossum ) and grapefems ( Botrychium ). This group has strong leaf dimorphism with large sporangia producing great quantities of spores (eusporangiate) in “fertile spikes” or “segments.” The sporangia lack a dehiscence mechanism (annulus) and the spores develop into underground gametophytes. The order Polypodiales has a diversity of growth habits and has small, specialized sporangia that produce smaller quantities of spores (leptosporangiate). Most species have sporangia with a definite dehiscence mechanism (annulus) with spores that usually produce green, above-ground gametophytes. The following families of this order, treated in this paper, are Aspleniaceae (spleenworts), Azollaceae (mosquito ferns), Blechnaceae (chain ferns), and Dennstaedtiaceae (bracken and hay-scented ferns). Information on specific and infraspecific taxa is set up in the following format: Number. Name author(s) [derivation of specific and infraspecific epithets]. VERNACULAR NAME. Habit; nativity (if exotic). Sporulating dates. Habitat data; highland provinces; relative abundance; [occurrence on Coastal Plain], Conservation status. Wetland indicator status. Comments. Synonyms. 39 Pteridophytes of NE Alabama Introduced taxa are followed by a dagger (t). Species of conservation concern are followed by a star (☆). The coded state ranks (ANHP 1994, 1996, 1997, 1999) are defined in Table 1. Wetland indicator status codes (Reed 1988) are defined in Table 2. Relative abundance is for occurrence in the study area and not for the whole state. Frequency of occurrence is defined as follows, ranging in descending order: common (occurring in abundance throughout), frequent (occurring throughout but not abundant), occasional (known in more than 50% of the region but in scattered localities), infrequent (known in less than 50% of the region in scattered localities or occurring in restricted habitats), rare (known from only a few counties and restricted to specific localities), and very rare (known from only a single or few populations; mostly narrow endemics, disjuncts, and peripheral taxa). Synonyms are from Mohr (1901)-- M; Small (1938)— S; Radford et al. (1968)— R; and Lellinger (1985) — L. Suggested pronunciation, author(s), date of citation, common name, and derivations are provided after each genus. Distribution maps are typically for 18 counties in the northeast region of Alabama. The maps are expanded to adjacent highland counties for taxa that are rare or peripheral. Key to symbols are as follows: Filled circle ( •) = documented at Jacksonville State University herbarium; filled square ( ■ ) = documented at another herbarium; open circle (O) = reported in literature. Table 1. Definition of state ranks. Code Definition 51 Critically imperiled in Alabama because of extreme rarity or because of some factor(s) making it especially vulnerable to extirpation from Alabama. 52 Imperiled in Alabama because of rarity or because of some factor(s) making it very vulnerable to extirpation from the state. 53 Rare or uncommon in Alabama. 54 Apparently secure in Alabama, with many occurrences. 55 Demonstrably secure in Alabama and essentially “ineradicable” under present conditions. SH Of historical occurrence , perhaps not verified in the past 20 years, and suspected to be still extant. SR Reported , but without persuasive documentation which would provide a basis for either accepting or rejecting the report. SU Possibly in peril in Alabama, but status uncertain. S? Not ranked to date. 40 Spaulding, et al. Table 2. Definition of wetland indicator codes. Code Status Probability of Occurrence OBL Obligate Wetland Species Occurs with estimated 99% probability in wetlands. FACW Facultative Wetland Species Estimated 67%-99% probability of occurrence in wetlands, 1%- 33% probability in nonwetlands. FAC Facultative Species Equally likely to occur in wetlands and nonwetlands(34%-66% probability). FACU Facultative Upland Species Estimated 67%-99% probability of occurrence in nonwetlands, l%-33% probability in wetlands. UPL Obligate Upland Species Occurs with estimated 99% probability in uplands. NI No Indicator Status Insufficient information available to determine an indicator status. Note: Positive or negative signs indicate a frequency toward higher (+) or lower (-) frequency of occurrence within a category. DIVISION III. POLYPODIOPHYTA Class 1. POLYPODIOPSIDA (=Filices) Order 1 . OPHIOGLOSSALES 1 . OPHIOGLOSSACEAE (Adder’s-tongue Family)* Contributed in part by Terri L. Ballard Selected reference: Wagner, W. H., Jr. and F. S. Wagner. 1993. Ophioglossaceae. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 85-106. 1. Sterile leaf blades pinnately dissected; veins free; fertile stalk with pinnately branched (grape-like) sporangia! clusters . Botrychium 1. Sterile leaf blades simple, unlobed; veins rejoining after branching (netted); fertile stalk with unbranched sporangial clusters . Ophioglossum 41 Pteridophytes of NE Alabama 1. BOTRYCHIUM {bo-TRIK-ee-um} Swartz 1800 • Grape-ferns • [Latin botry, bunch (of grapes), and -oides, like; in reference to the sporangial clusters.] Selected reference: Wagner, Jr. W. H. and F. S. Wagner. 1983. Genus communities as a systematic tool in the study of New World Botrychium (Ophioglossaceae). Taxon 32: 51-63. 1 . Sterile leaf blades thin, herbaceous, and absent during winter; fertile stalk arising from base of sterile blade; common stalk raised well above ground; leaf sheaths open . B. virginianum 1. Sterile leaf blades leathery, evergreen or herbaceous, but present through winter; fertile stalk not arising at base of sterile leaf blade; common stalk at or near ground; leaf sheaths closed. 2. Leaves sessile or short-stalked, often prostrate on ground; plants ususally with 2 or more leaves; leaflets as long as wide (fan-shaped) and lacking a central vein; fertile stalk (sporophore) producing spores in late winter or early spring; new leaves appear in late fall and die in spring; roots yellowish and smooth . B. lunarioides 2. Leaves long-stalked (more than 15 mm long) and erect or ascending; plants with 1 or more leaves; leaflets usually longer than wide, with a weak to strong central vein; fertile stalk producing spores in summer or fall; new leaves appear in spring or summer and last until the following spring; roots brownish and ribbed. 3. Leaflets (divisions of sterile blade) not much longer than broad, tips obtuse; sterile leaf blades often 2 per season, usually sprawling; leaflets with weak central vein . B. j&nmanii 3. Leaflets much longer than broad, tips acute; sterile leaf blades normally 1 per year, mostly upright; leaflets with strong central vein. 4. Sterile leaf blades 2-pinnate; small lateral leaflets (pinnae or pinnules) oblong and somewhat rounded; leaves usually green in winter . B. biternatum 4. Sterile leaf blades 3-pinnate; small lateral leaflets (pinnae or pinnules) rhomboidal and angular; leaves usually bronze in winter . B. dissection 1. Botrychium biternatum (Savigny) Underwood [twice ternate], SOUTHERN GRAPEFERN; SPARSE-LOBED GRAPEFERN. Figure 1. Evergreen perennial. Sporulates August - October. Woods, stream banks, and fields; all highland provinces; frequent; [Coastal Plain], Wetland Indicator Status, FAC. The specific epithet is referring to the sterile blades that are divided into three major parts, each of which is again divided into three parts (Theiret 1980). 2. Botrychium dissectum Sprengel [dissected], DISSECTED GRAPEFERN; COMMON GRAPEFERN. Figure 2. Evergreen perennial. Sporulates August - October. Alluvial woods and stream banks; all highland provinces; frequent; [Coastal Plain]. Wetland Indicator Status, FAC. Leaves persist throughout the winter, darkening to bronze or copper. The highly dissected form of this variable species (f. dissectum) is much less frequent in northeast Alabama than the less dissected form (f. obliquum). Synonym: Botrychium obliquum (Muhlenberg) Small— M,S. 3. Botrychium jenmanii ☆ Underwood [George Jenman (1845-1902), British botanist], ALABAMA GRAPEFERN; JENMAN’S GRAPEFERN. Figure 3. Evergreen perennial. Sporulates August - October. Open grassy sites and low pine woodlands; Cumberland Plateau and Piedmont; very rare; [Coastal Plain]. State Rank, SH. Wetland Indicator Status, NI. This fern 42 Spaulding, et al. was once listed as imperiled (S2) in the state, but now listed as of historical occurrence because it had not been seen in awhile. In spring of 2001, Brian Keener and Dan Spaulding collected it in a graveyard in Wadley, Alabama. William Maxon discovered this fern near Mobile, Alabama in 1906, but it was actually first collected from the Piedmont of Georgia in 1900 (Dean 1969). It is similar to B. lunarioides and often associated with it in lawns of cemeteries. Botrychium jenmanii is thought to be a fertile hybrid between B. lunarioides and B. biternatum. Synonym: Botrychium alabamense Maxon— S.R. 3. Botrychium lunarioides fr Michaux [like B. lunaria, moonwort], WINTER GRAPEFERN; PROSTRATE GRAPEFERN. Figure 4. Deciduous perennial (leaves dying by April). Sporulates January - April. Dry soil in grassy areas such as cemeteries, pastures, and mown fields; Cumberland Plateau and Piedmont; rare; [chiefly Coastal Plain], State Rank, SH. Wetland Indicator Status, NI. This fern should probably be listed as SI or S2 because it was recently collected in Morgan County by Alvin Diamond and in Randoph County by Brain Keener and Dan Spaulding. It is easily overlooked, and you have to often crawl on the ground to find it. Botrychium lunarioides is often associated with another inconspicuous family member, Ophioglossum crotalophoroides (bulbous adder’s-tongue). 4. Botrychium virginianum (Linnaeus) Swartz [Virginian]. RATTLESNAKE FERN; VIRGINIA GRAPEFERN. Figure 5. Deciduous perennial. Sporulates April - June. Woods, thickets, and meadows; all highland provinces; common; [Coastal Plain], Wetland Indicator Status, FAC+. This species is the most widespread Botrychium of North America (Wagner and Wagner 1993). The name “rattlesnake fern” is in reference to the fertile stalk which supposedly resembles the tail of this snake (Clute 1938). Indians used the rootstock and made a poultice for snakebites, bruises, cuts, and sores (Foster and Duke 1990). Synonym: Osmundopteris virginiana (Linnaeus) Small— S. 2. OPHIOGLOSSUM {oh-fee-oh-GLOSS-um} Linnaeus 1753 • Adder’s-tongue Ferns • [Greek ophis, snake, and glossa, tongue, referring to the sporulating portion of the fertile rachis.] 1. Rhizome (underground stem) globose-bulbous; leaf blades debate to cordate . . O. crotalophoroides 1. Rhizome cylindrical; leaves ovate to elliptical. 2. Apex of sterile leaf blades ending abruptly in a small slender point; veins of sterile leaf blades forming smaller secondary areoles (“circles”) within larger primary areoles; fresh plants malodorous; plants of limestone outcroppings . O. engelmannii 2. Apex of sterile leaf blades rounded; veins of sterile blades forming areoles without secondary areoles, but with included free vienlets; fresh plants not malodorous; plants not usually associated with limestone outcroppings . O. vulgatum Note: Dr. Herb Wagner (1999) states that Ophioglossum nudicaule Linnaeus /, Slender Adder’s-tongue, and Ophioglossum petiolatum Hooker, Stalked Adder’s-tongue, are likely to be found in habitats similar to those of O. crotalophoroides, but have not been documented from our study area. These plants are easily over-looked because of their smaller size. Both adder’ s-tongues are more common on the coastal plain. Like O. crotalophoroides, they usually have two to three leaves per stem, but lack a corm-like 43 Pteridophytes of NE Alabama rhizome and cordate-deltate leaves. Ophioglossum midicaule is less than 6 cm tall with leaf blades arising near base of plant and O. petiolatum is more than 6 cm tall with leaf blades arising near the middle of plant. The latter species is probably not native and was introduced through plant nurseries (Nelson 2000). 1. Ophioglossum crotalophoroides vrWalter [like a rattle-bearer], BULBOUS ADDER'S-TONGUE. Figure 6. Deciduous perennial. Sporulates February - September. Open grassy areas such as lawns, fields, and cemeteries; Cumberland Plateau; rare; [chiefly Coastal Plain], State Rank, previously S3 (ANFIP 1994). Wetland Indicator Status, FAC-. First discovered in the late 18th century by Thomas Walter who included it in Flora Caroliniana in 1788 (Dean 1969). The specific epithet is referring to the sporangia resembling the rattles of a rattlesnake ( Crotalus ). This species is easily overlooked, it is best to get down on your knees with your head close to the ground in order to find it. 2. Ophioglossum engelmannii ☆ Prantl [G. Engelmann, 1809-1884], LIMESTONE ADDER'S- TONGUE. Figure 7. Deciduous perennial. Sporulates March - June. Mostly in soil over limestone in open fields, pastures, and cedar glades; Interior Low Plateau, Cumberland Plateau; infrequent; [Coastal Plain], State Rank, S2S3. Wetland Indicator Status, FACU. This species forms extensive colonies by vegetative reproduction. Karl Prantl (1849-1893), a German botanist, named this species in 1883 in honor of George Engelmann (1809-1884), an American plant taxonomist, (Nelson 2000). 3. Ophioglossum vulgaium Linnaeus [common], SOUTHERN or COMMON ADDER'S-TONGUE. Figure 8. Deciduous perennial. Sporulates April - July. Low woods, floodplains, and meadows; all highland provinces; occasional; [Coastal Plain]. Wetland Indicator Status, FACW. Synonyms: Ophioglossum vulgatum var. pycnostichum Femald— R; Ophioglossum pycnostichum (Femald) Love & Love-- L. Order 2. POLYPODIALES (=Filicales) 1 . ASPLENIACEAE (Spleenwort Family) 1. ASPLENIUM {ess-PLEEN-ee-um} Linnaeus 1753 • Spleenworts • [Greek splen, for spleen; once thought useful for diseases of the spleen.] Selected reference: Wagner, W. H. Jr., R. C. Moran, and C. R. Werth. 1993. Aspleniaceae. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 228-245. 1. Leaves simple or deeply lobed, not divided to midvein; leaf apices (tips) usually long tapering (acute to acuminate in A. scolopendrium). 2. Leaf blades deeply lobed . A. pinnatifidum 2. Leaf blades simple, not lobed. 3. Leaf apex long tapering (attenuate) and often rooting at tip; veins forming areoles 44 Spaulding, et al. (anastomosing); sori single along veins . A. rhizophyllum 3. Leaf apex acute to acuminate and not rooting at tips; veins free; sori in pairs along veins . A. scolopendrium 1. Leaves pinnately divided; leaf tips usually acute (rarely rounded or acuminate), not long tapering. 4. Leaf blades 1 -pinnate and leaflets (pinnae) not lobed. 5. Leaflets mostly alternate; base of leaflets with auricles (lobes) that usually overlap rachis . A. platyneuron 5. Leaflets mostly opposite; base of leaflets not overlapping rachis. 6. Leaflets strongly asymmetrical, midvein running along basal edge of blade; sori few (usually 1) and located on only one side of main vein (costa) . . A. monanthes 6. Leaflets symmetrical, midvein running along the middle of blade; sori numerous (more than 4) and located on both sides of the main vein. 7. Leaflets oblong (longer than wide) and usually more than 9 mm long; auricles (lobes) present at base of leaflets . A. resiliens 7. Leaflets mostly oval (almost as long as wide) and usually less than 9 mm long; auricles lacking at base of leaflets . A. trichomanes 4. Leaf blades 2 to 4-pinnate or pinnae (leaflets) deeply lobed (at least lower ones). 8. Blades with 2-5 pairs of pinnae; leaf stalks (petioles) entirely green; plants on calcareous (limestone) rock . A. ruta-muraria 8. Blades with more than 5 pairs of pinnae; leaf stalks darkened, at least at base; plants on acidic (usually sandstone) rock. 9. Leaf stalks (petioles) dark colored only at base; most pinnae divided . . A. montanum 9. Leaf stalks and lower portion of rachis darkly colored; only the lower pinnae divided (bipinnate) . A. bradleyi Note: The following hybrids are known to occur in our study area, but are rare (figure 9). (1) Asplenium x gravesii Maxon, Grave’s Spleenwort, is an infertile hybrid between A. pinnatifidum and A. bradleyi. The frond (leaf blade) of this hybrid is pinnate except for the apex which is usually deeply lobed (pinnatifid). Its leaf stalk (petiole) and portions of rachis are dark colored. Maxon named this hybrid in honor of E. W. Graves, who according to Dean (1969) discovered it in Alabama in 1917. However, Short (1999) states that Dean was mistaken and that the type locality is actually in Dade County, Georgia. (2) Asplenium x trudellii Wherry, Trudell’s Spleenwort, is an infertile hybrid between A. montanum and A. pinnatifidum. The frond of this hybrid is often deeply lobed (pinnatifid) near the apex and pinnate to pinnate-pinnatifid at the base. Its leaf stalk (petiole) is mostly green and it occurs on rock, like its parents. (3) Asplenium x ebenoides R. R. Scott, Scott’s Spleenwort, an infertile hybrid (in our range) between A. rhizophyllum and A. platyneuron. One population in Hale County (southern Alabama), has doubled its chromosomes and has become a fertile allotetraploid. This fertile population was discovered in 1 874 by Julia L. Tutwiler, a 45 Pteridophytes of NE Alabama teacher near Havana, Alabama (Walter et al. 1982). This hybrid was named by Robert R. Scott who discovered it near Philadelphia in 1862 (Dean 1969). The frond is usually deeply lobed (pinnatifid) with a long tapering tip, though it is sometimes pinnate at the very base. The lobes of the leaf blade are uneven in length, giving it an irregular outline and its leaf stalk (petiole) is dark throughout its length. 1. Asplenium bradleyi fr D. C. Eaton [F. H. Bradley, 1838-1879]. CLIFF or BRADLEY’S SPLEENWORT. Figure 10. Evergreen perennial. Sporulates April - October. Acidic rock cliffs and ledges (usually composed of sandstone) in shady, dry-mesic environments; Cumberland Plateau, Ridge and Valley, upper Piedmont; infrequent. State Rank, S2. Wetland Indicator Status, UPL. This species is a fertile hybrid (allotetraploid) between A. montanum and A. platyneuron (Weakly 1997). This fern was discovered in Tennessee by Frank Bradley in 1870 (Dean 1969). 2. Asplenium monanthes ☆ Linnaeus [one- “flowered”]. SlNGLE-SORUS SPLEENWORT. Figure 1 1. Evergreen perennial. Sporulates April - October. Growing in crevices on rock in subacid to circumneutral soil; mossy granite-schist ledges in the upper Piedmont and vertical limestone sinks in the Cumberland Plateau; very rare. State Rank, S 1 . Wetland Indicator Status, NI. This fern was not discovered in the Southeastern U.S. until 1946 in South Carolina. Spores from Mexico are thought to have been brought in by a hurricane (Wherry 1972). 3. Asplenium montanum Willdenow [montane]. MOUNTAIN SPLEENWORT. Figure 12. Evergreen perennial. Sporulates May - October. Sandstone (acidic) rock outcrops and bluffs; Interior Low Plateau, Cumberland Plateau, Ridge and Valley, upper Piedmont; infrequent. Wetland Indicator Status, UPL. 4. Asplenium pinnatifidum Nuttall [pinnatifid], LOBED SPLEENWORT. Figure 13. Evergreen perennial. Sporulates May - October. Rock outcrops in gorges, usually on sandstone; acidic soils in forests; Cumberland Plateau; infrequent. Wetland Indicator Status, UPL. 5. Asplenium platyneuron (Linnaeus) Britton, Stems, & Poggenburg [broad-nerved], EBONY SPLEENWORT. Figure 14. Evergreen perennial. Sporulates April - November. Forests, woodlands, borders of wood, thickets, rock outcroppings, disturbed sites, usually in shallow, dry- mesic soils, acidic to circumneutral substrates; all highland provinces; common; [Coastal Plain]. Wetland Indicator Status, FACU. The specific epithet is a misnomer based on an early drawing with a large, exaggerated rachis (Nelson 2000). Synonym: Asplenium platyneuron (Linnaeus) Oakes— M, S, R. 6. Asplenium resiliens Kunze [springing back], BLACK- STEMMED SPLEENWORT. Figure 15. Evergreen perennial. Sporulates April - November. Rock outcrops, circumneutral soils, frequently on limestone; Interior Low Plateau, Cumberland Plateau, Ridge and Valley, upper Piedmont; frequent; [Coastal Plain], Wetland Indicator Status, UPL. The specific epithet may refer to the flexible leaf stalks (petioles). Synonym Asplenium parvulum Martens & Galeotti— M. 7. Asplenium rhizophyllum Linnaeus [rooting leaf], WALKING FERN. Figure 16. Evergreen perennial. Sporulates May B October. Shaded rock ledges and boulders, frequently on limestone though occasionally on acidic rocks; Interior Low Plateau, Cumberland Plateau, Ridge and Valley; occasional; [rarely Coastal Plain], Wetland Indicator Status, UPL. Synonym: Camptosorus rhizophyllus (Linnaeus) Link— M. 46 Spaulding, et al. 8. Asplenium ruta-muraria ☆ Linnaeus [rue - growing on walls]. WALL-RUE SPLEENWORT. Figure 17. Evergreen perennial. Sporulates May - October. Crevices of limestone cliffs and ledges; Interior Low Plateau, Cumberland Plateau, Ridge and Valley; rare. State Rank, S2. Wetland Indicator Status, UPL. Some authors consider our species as var. cryptolepis (Fernald) Wherry, which only differs slightly from the European species. In Europe this fern is common on masonry walls. Synonym: Asplenium cryptolepis Fernald— S. 9. Asplenium scolopendrium ☆ Linnaeus [Greek name of a similar plant] var. americanum (Fernald) Kartesz & Gandhi [American], HART’S-TONGUE FERN. Figure 18. Evergreen perennial. Sporulates September - December. Deeply shaded limestone sinks with cool, humid air; Cumberland Plateau; very rare. Federal Status, Threatened; State Rank, SI. Wetland Indicator Status, NI. Although difficult to locate the habitat, various spelunkers have actively sought such caves and explored them with the intent of locating new populations. It was discovered growing in Alabama in 1978 during a caving expedition in Jackson County (Short 1979). First described by Linnaeus in 1753 from European plants. It was first discovered in North America in 1807 by Frederick Pursh, near Syracuse, New York (Clute 1938). Other vernacular names for this fern include Hound’s-tongue and Seaweed Fern, all which allude to the shape of the fronds. Synonyms: Phyllitis scolopendrium (Linnaeus) Newman var. americana Fernald— L. 10. Asplenium trichomanes ir Linnaeus [hair cup]. MAIDENHAIR SPLEENWORT. Figure 19. Evergreen perennial. Sporulates May - November. Rocky slopes and bluffs, frequently growing on rocks; Interior Low Plateau, Cumberland Plateau, Ridge and Valley, upper Piedmont; infrequent. State Rank, S2S3. Wetland Indicator Status, UPL. Two infraspecific taxa, subsp. trichomanes and subsp. quadrivalens D. E. Meyer, occur in North America. Only the type species is found in our state. Asplenium trichomanes subsp. trichomanes also occurs in Europe. 2. AZOLLACEAE (Mosquito Fern Family) 1. AZOLLA {ah-ZOH-la} Lamarck 1783 • Mosquito Ferns • [Greek azo, to dry, and ollyo, to kill; alluding to the possibility of drought killing this aquatic fern.] Selected references: Lumpkin, T. A. 1993. Azollaceae. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 338-339. Svenson, H. K. 1944. The New World species of Azolla. Amer. Fern J. 34: 69-84. 1. Azolla caroliniana Willdenow [Carolinian]. MOSQUITO FERN; WATER FERN. Figure 20. Evergreen, floating aquatic. Rarely collected with sporocarps (hard, pea-shaped bodies that contain spores). Stagnant or slow-moving water of lakes, ponds, marshes, and streams, though may be stranded on wet muck when pool level drops; along the Tennessee River Valley in the Interior Low Plateau and Cumberland Plateau; infrequent; [chiefly Coastal Plain]. Wetland Indicator Status, OBL. Economically important because of its symbiotic relationship with the nitrogen-fixing blue-green alga, Anabaena azollae. Used as a “green” fertilizer with crops such 47 Pteridophytes of NE Alabama as rice or a nutritional supplement when mixed with livestock feed (Dunbar 1989). Plants turn red when under stress from factors such as high temperatures or poor nutrition. Growth of this fern can be so dense over the water surface that it can exclude mosquito larvae, hence the common name (Snyder and Bruce 1986). Short (1999) believes this is essentially a Coastal Plain plant. It was probably introduced into the Tennessee Valley and it grows there today because the Tennessee Valley Authority (TVA) lakes provide the still-water type of habitat it needs. 3. BLECHNACEAE (Chain Fern Family) Selected Reference: Cranfill, R. B. 1993. Blechnaceae. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 223-224. 1. WOODWARDIA {wood-WAR-dee-uh} Smith 1793 • Chain Ferns • [in lionor of English botanist, Thomas Jenkinson Woodward, 1745-1820.] The common name for this genus is in reference to the “chain-lil^e” appearance of the sori. Recent research suggests that the two local elements represent two genera: Lorinseria and Anchistea (Wagner 1999). Selected Reference: Cranfill, R. B. 1993. Woodwardia. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 226-227. 1. Leaves dimorphic; fertile leaves pinnate, sterile leaves mostly pinnatifid (deeply lobed); veins conspicuously netted . W. areolata 1. Leaves monomorphic (sterile and fertile leaves similar); leaves pinnate with pinnatifid leaflets; veins mostly free . W. virginica 1. Woodwardia areolata (Linnaeus) T. Moore [with areoles]. NETTED or NET-LEAVED CHAIN Fern. Figure 21. Deciduous perennial. Sporulates late April - November. Swamps, stream banks, bogs, seepage slopes, and wet mixed woods; all highland provinces; frequent; [Coastal Plain], Wetland Indicator Status, OBL. The common name refers to the noticeably netted venation of the sterile leaves. This species is often confused with Sensitive Fern ( Onoclea sensibilis). Sterile leaves of W. areolata usually have an alternate pinna arrangement and finely serrate leaf margins; whereas, O. sensibilis usually has an opposite pinna arrangement and entire leaf margins. Synonym: Lorinseria areolata (Linnaeus) Presley— S. 2. Woodwardia virginica (Linnaeus) Smith [Virginian], VIRGINIA or SOUTHERN CHAIN FERN. Figure 22. Deciduous perennial. Sporulates July - September. Creek banks, open wet areas, and forested wetlands; Cumberland Plateau, Ridge and Valley; infrequent; [chiefly Coastal Plain], Wetland Indicator Status, OBL. This fern is used by the Chain Fern Borer Moth ( Papaipema stenocelis ) as a food source (Dunbar 1989). This species resembles Cinnamon Fern ( Osmunda cinnamomea), but Woodwardia virginica can be distinguished having the following suite of characteristics: blackish leaf stalk , absence of woolly, brown hairs, and non-clustered (non¬ circular) growth pattern. Synonym: Anchistea virginica (Linnaeus) Presley— S. 48 Spaulding, et al. 4. DENNSTAEDTIACEAE (Cuplet Fern Family)* Contributed in part by Timothy L. Hofmann Selected reference: Cranfill, R. B. 1993. Dennstaedtiaceae. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 198B205. 1. Leaf axis 3-branched; leaf blades triangular and lacking glands (eglandular); sori continuous and covered by rolled under (revolute) leaflet margin . Pteridium 1. Leaf axis not branched; leaf blades broadly elliptic (not triangular) and glandular; sori distinct and not covered by leaflet margin . Dennstaedtia Note: Hypolepis repens (Linnaeus) C. Presl, Bramble Fern, sometimes escapes from cultivation and is native to tropical America. An old record for this species was collected in Jefferson County. It is similar to Hay-scented Fern ( Dennstaedtia punctilobula), but leaves are 4-pinnate-pinnatifid (instead of 2-pinnate-pinnatifid in Dennstaedtia) and has “flaps” (false outer indusia) that cover the sori along margins of the leaflets. 1. DENNSTAEDTIA {den-STET-ee-uh} Bemhardi 1802 • Cuplet Ferns • [Named for August W. Dennstaedt, 1776-1826, a German botanist.] Selected references: Nauman, C. E. and A. Murray Evans. 1993. Dennstaedtia. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 19-201. Tyron, R. M., Jr. 1960. A review of the genus Dennstaedtia in America. Contr. Gray Herb. 187: 23-52. 1. Dennstaedtia punctilobula ☆ (Michaux) T. Moore [with dotted lobes], HAY-SCENTED FERN; BOULDER Fern. Figure 23. Deciduous perennial. Sporulates June - September. Shaded sandstone bluffs and rocky slopes; Cumberland Plateau, Ridge and Valley, Upper Piedmont; infrequent. State Rank, previously S3 (ANHP 1994). Wetland Indicator Status, UPL. Fronds have the scent of newly mown hay, hence the common name. The name “boulder fern” comes from the common association of this fern with rocky habitats (Clute 1938). Cherokee Indians made a tea from the plant to help control chills (Dunbar 1989). 2. PTERIDIUM {ter-RlD-ee-um} Gleditschex Scopoli 1760 • Bracken Ferns • [Greek, pteridion, diminutive of Pteris, a wing; an ancient name for a fern.] “Bracken” or “Brake” is from an old Saxon word meaning fallow or clearing, alluding to habitats of this fern. Selected reference: Jacobs, C. A. and J. H. Peck. 1993. Pteridium. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 201-204. 49 Pteridophytes of NE Alabama 1 . Terminal pinnules (leaflets) 6 to 1 5 times longer than wide, undersurface of leaflets (abaxial axes) remotely pilose to glabrous . P. aquilinum var. pseudocaudatum 1. Terminal pinnules 2 to 4 times longer than wide, undersurface of leaflets (abaxial axes) villous . . . P. aquilinum var. latiusculum 1. Pteridium aquilinum (Linnaeus) Kuhn var. latiusculum (Desvaux) Underwood [broad], EASTERN Bracken Fern; Pasture Brake. Figure 24. Deciduous perennial. Sporulates July - September. Open woods, roadsides, pastures, meadows, usually in acid to strongly acid soils; Cumberland Plateau; rare. Wetland Indicator Status, FACU. The species has a world-wide distribution. Ashes of bracken fern were used to make soap because of their high potash content (Clute 1938). Bracken Fern is the host plant for the Bracken Borer Moth, ( Papaipema pterisii). Synonyms: P ter is latiuscula Desvaux— S. 2. Pteridium aquilinum (Linnaeus) Kuhn [of an eagle] var. pseudocaudatum (Clute) Heller [false tail]. TAILED BRACKEN FERN; SOUTHERN BRACKEN. Figure 25. Deciduous perennial. Sporulates July - September. Forms large colonies, in barrens, open woods, in sandy, acid soils; all highland provinces; common; [Coastal Plain], Wetland Indicator Status, FACU. Deep rhizomes are protected from fires. Fiddleheads were eaten as a vegetable, but recent studies have shown them to be carcinogenic. Plants contain the enzyme thiaminase and if consumed in large quantities by livestock may cause poisonings (Dunbar 1989). The specific epithet alludes to the wing-shaped fronds or possibly to the fiddleheads resemblance to an eagle’s foot. Synonym: Pteris aquilina Linnaeus var. pseudocaudata Clute— M. REFERENCES CITED Alabama Natural Heritage Program [ANHP], 1994. Vascular Plant Inventory Tracking List, April edition. Montgomery, Alabama. Alabama Natural Heritage Program. 1996. Species Inventory List. Montgomery, Alabama. Alabama Natural Heritage Program. 1997. Inventory List of Rare, Threatened, and Endangered Plants, Animals, and Natural Communities of Alabama. Montgomery, Alabama. Alabama Natural Heritage Program. 1999. Inventory List of Rare, Threatened, and Endangered Plants, Animals, and Natural Communities of Alabama. Montgomery, Alabama. Clute, W. N. 1938. Our Ferns: Their Haunts, Habits and Folklore, 2nd ed. Frederick A. Stokes Company, New York. Dean, B. E. 1969. Ferns of Alabama. Southern University Press, Birmingham. Dunbar, Lin. 1989. Ferns of the Coastal Plain, their Lore, Legends and Uses. University of South Carolina Press. Foster, S. and J. A. Duke. 1990. A Field Guide to Medicinal Plants, Peterson Field Guide Series. Houghton Mifflin Co., Boston. Lellinger, D. B. 1985. A Field Manual of the Ferns and Fern-allies of the United States and Canada. Washington, D. C. Mohr, C. 1901. Plant Life of Alabama. Cont. U. S. Nat. Herb. 6. Nelson, G. 2000. Ferns of Florida. Pineapple Press, Sarasota. Radford, A. E., H. E. Ahles, and C. R. Bell. 1968. Manual of the Vascular Flora of the 50 Spaulding, et al. Carolinas. The University of North Carolina Press. Reed, P.B. 1988. National List of Plant Species that Occur in Wetlands: Southeast (Region 2). U.S. Fish and Wildlife Service, Biological Rep. 88 (24), 244 p. Short, J. W. 1979. Phyllitis scolopendrium newly discovered in Alabama. Amer. Fern J. 69: 47B48. Short, J. W. 1999. Personal letter to Dan Spaulding at the Anniston Museum of Natural History, 21 June. Small, J. K. 1938. Ferns of Southeastern States. Published by author. New York. Snyder, L. H. and J. G. Bruce. 1986. Field Guide to the Ferns and Other Pteridophytes of Georgia. The University of Georgia Press. Theiret, J. W. 1980. Louisiana Ferns and Fern Allies. Lafayette Natural History Museum in conjunction with the University of Southwest Louisiana. Lafayette, Louisiana. Wagner, W. H, 1999. Personal letter and notes to Daniel D. Spaulding at the Anniston Museum of Natural History, 16 November. Wagner, W. H., Jr. and F. S. Wagner. 1993. Ophioglossaceae. In: Flora of North America Editorial Committee, eds. 1993+. Flora of North America North of Mexico. 3+ vols. New York and Oxford. Vol. 2, pp. 85B106. Walter, K. S., W. H. Wagner, Jr., and F. S. Wagner. 1982. Ecological, biosystematic, and nomenclatural notes on Scott’s Spleenwort, X Asplenosorus ebenoides. Amer. Fern J. 72: 65B75. Weakly, A. S. 1997. Flora of the Carolinas and Virginia, working draft. The Nature Conservancy, Regional Office, Southern Conservation Science Department. 51 Pteridophytes of NE Alabama Figure 1. Botrychium biternatum- Southern Grapefern Figure 2. Botrychium dissectum- Dissected Grapefern 52 Spaulding, et al. Figure 3. Botrychium jenmanii- Alabama Grapefern Figure 4. Botrychium lunarioides- Winter Grapefern 53 Pteridophytes of NE Alabama Figure 5. Botrychium virginianum- Rattlesnake Fern Figure 6. Ophioglossum crotalophoroides- Bulbous Adder's-tongue 54 Spaulding, et al. Figure 7. Ophioglossum engelmannii - Limestone Adder's-tongue Figure 8. Ophioglossum vulgatum- Common Adder's-tongue 55 Pteridophytes of NE Alabama A. x gravesii- Grave's Spleenwort A. x ebenoides- Scott's Spleenwort Figure 9. Asplenium hybrids 56 Spaulding, et al. / Figure 10. Asplenium bradleyi- Cliff Spleenwort Figure 11. Asplenium monanthes- Single-sorus Spleenwort 57 Pteridophytes of NE Alabama Figure 12. Asplenium montanum- Mountain Spleenwort Figure 13. Asplenium pinnatifidum- Lobed Spleenwort 58 Spaulding, et al. Figure 15. Asplenium resiliens- Black-stemmed Spleenwort Figure 14. Asplenium platyneuron- Ebony Spleenwort 59 Pteridophytes of NE Alabama Figure. 16. Asplenium rhizophyllum- Walking Fern Figure 17. Asplenium ruta-muraria- Wall-rue Spleenwort 60 Spaulding, et al. Asplenium scolopendrium Figure 18. Asplenium scolopendrium- Hart's-tongue Fern 4. Figure 19. Asplenium tricnomanes- Maiaennair Spleenwort 61 Pteridophytes of NE Alabama Figure 20. Azolla caroliniana- Mosquito Fern Figure 21. Woodwardia areolata- Netted Chain Fern 62 Spaulding, et al. Figure 22. Woodwardia virginica- Virginia Chain Fern 63 Pteridophytes of NE Alabama Figure 24. Pteridium aquilinum var. latiusculum- Eastern Bracken Fern Figure 25. Pteridium aquilinum var. pseudocaudatum- Tailed Bracken Fern 64 Journal of the Alabama Academy of Science, Vol. 72, No. 1 , January 200 1 . THE VASCULAR FLORA OF DALE COUNTY LAKE, ALABAMA Michael Woods, Brian Prazinko Alvin R. Diamond, Jr. Department of Biological and Environmental Sciences Troy State University Troy, Alabama 36082 ABSTRACT A survey of the vascular flora of the area surrounding Dale County Lake was conducted from May 1998 through August 2000. The study site, located in the central section of Dale County, was comprised of 106 land ha and 37 water ha. Five major habitats, shoreline, mesic woods, xeric woods, wetland, and grassy area were found to occur at the study site. Collections were made once weekly from April through November and twice monthly from December through March. Similarity indices were determined by comparing this study with two other recent floristic studies from south Alabama. The 407 species representing 265 genera and 98 families found to occur at Dale County Lake are presented in an annotated checklist. INTRODUCTION Publications dealing with the flora of Alabama are rare and are becoming increasingly outdated. Comprehensive treatments are limited to Mohr (1901) and Small (1933), while Harper (1943), Godfrey & Wooten (1979, 1981) and Godfrey (1988) concentrate on specific plant communities. In their work on the fauna and flora of Fort Rucker, Mount and Diamond (1992) collected 549 taxa of vascular plants. Diamond and Freeman (1993) reported 908 taxa of vascular plants from Conecuh County. Crouch and Golden (1997) listed 450 species of vascular plants from the area along the Tombigbee River in northeastern Choctaw County. Woods and Reiss (1998) reported 274 species and varieties from the area around Pike County Lake. In the most recent study, Rundell and Woods (2001) reported 507 species and varieties from the area around Ech Lake in the west-central section of Dale County, on the Fort Rucker Army Installation. Although these studies were comprehensive treatments of their collection area, additional records are needed to adequately represent the diversity of plants found in south Alabama. DESCRIPTION OF STUDY AREA The Dale County Lake study area is located in the central part of Dale County, Alabama (31°45'N, 85°64'W) immediately outside and north-northeast of the city limits of Ozark. This places the study area entirely in the eastern division of the Southern Red Hills of 65 Flora of Dale County Lake the Coastal Plain Province. The soils are characteristic of those formed in the humid, subtropical climate of the southeastern section of the United States. They are derived from the sandy red clays of the Lafayette formation. The topsoil is paler and sandier, except where washed off on steep slopes (Harper 1943). The climate in Dale County is considered subtropical-humid with long hot summers and short, mild winters. The annual growing season is approximately 250 days. Summer temperatures of above 30°C occur frequently and temperatures above 40°C are not uncommon. Cumulatively, the climate is considered very mild and therefore favors the growth of a wide variety of plant life (Brown 1999). The Alabama Department of Conservation initiated the construction of Dale County Lake in 1956 and it was completed and opened to the public in 1958. The damming of Panther Creek, which serves as the primary inflow and outflow stream, formed the lake. Two smaller streams also serve as inflows for the lake. The study site comprised 106 ha of mostly forested, gently sloping land surrounding the 37 ha lake. The land and the lake are owned by the Alabama Department of Conservation and Natural Resources and managed by the City of Ozark Department of Parks and Recreation. DESCRIPTION OF HABITATS Five habitats were defined at Dale County Lake on the basis of topography, moisture content, and vegetation (Table 1). The shoreline habitat consisted of a narrow band of land along the edge of the lake. Found in this habitat were plants growing at the water edge, ranging from totally submersed to partially or completely immersed. The mesic woods habitat, which consisted of approximately 35 ha occurred on flat to gently rolling slopes that border the shoreline habitat. This area was composed of rich soils with an abundance of leaf litter. Soil moisture was abundant, and due to dense overhead vegetation light was limited at ground level. The xeric woods habitat, which consisted of approximately 50 ha consisted of well-drained sandy soils found at higher elevations on the study site. One exception was along the northeast side of the lake where the xeric habitat extended down to the shoreline. Loblolly Pines, which were planted more than twenty-five years ago, were the dominant trees in this area. Wetlands, which occurred in low-lying areas along Panther Creek and the two smaller inlet streams comprised approximately 12 ha of land. Most of this area only recently became wetlands due to the successful damming of the streams by beavers. In these wetlands, water was present at the soil surface throughout most of the year. In some areas, where the beavers were successful in damming the streams, water levels ranged from 0.5 to 1.0 meter deep throughout the year. Approximately 9 ha of grassland habitat were located around the central office, parking lot, boat-ramp, and land piers. Additional grassland habitat was found along the 5.6 km walking trail around the lake. Most of these grasslands were closely cut to benefit fishermen and those involved in other recreational activities. METHODS The systematic collection of vascular plants of the Dale County Lake study area began in May of 1998 and continued through August of 2000. To obtain a more complete floristic list the five specific habitats were collected twice weekly from April through November and twice monthly from December through March. All seed plants, with the exception of some 66 Woods, et al. trees were collected with either flower or fruit. Seedless vascular plants were collected with sporangia. Voucher specimens were deposited in the herbarium of Troy State University (TROY). Table 1 . Habitats and the number of species collected at each Habitats Number of Species Collected Number of Non-Native Species Shoreline 27 1 Mesic Woods 197 23 Xeric Woods 52 3 Wetland 58 5 Grassland 65 21 Floristic similarity between Dale County Lake and two other similar studies in South Alabama was determined using Sorensen's similarity coefficient (Mueller-Dobois and Ellenberg 1974) where: IS = 2C/A+B x 100 IS = Index of Similarity A = Total number of taxa at Ech Lake B = Total number of taxa at comparison study site C = Total number of taxa shared by A and B This analysis provided a means of determining statistical similarities between this study site and other floristic studies conducted from south Alabama. Sources used for identification and determination of non-native species were: Radford, Ahles and Bell (1968), Hitchcock (1971), Godfrey and Wooten (1979, 1981), Cronquist (1980), Godfrey (1988), Clewell (1985) and Isely (1990). RESULTS During the 28-month study period, 407 species representing 98 families and 265 genera were collected at Dale County Lake. The largest families were Asteraceae with 55 species, Poaceae with 34 and Fabaceae with 31. Carex and Smilax represented the largest genera with 7 species each. Cyperus, Eupatorium, and Quercus each had 6 species. Fifty- three introduced species were collected with at least one non-native species from each of the 67 Flora of Dale County Lake five habitats we characterized (Table 1). Murdannia keisak, an introduced species from Asia, was collected during this study. This was the second reported location of this species from Dale County and only the third report from Alabama (Rundell and Diamond 1999). The two floristic investigations used to determine the index of similarity were chosen because they share certain features with the present study, such as size of the study areas, physiographic factors, and climate. In the study “The vascular flora of Ech Lake, Alabama” (Rundell and Woods 2001), an area of 162 ha located in the west-central section of Dale County, a total of 507 species, representing 289 genera and 102 families were reported. Woods and Reiss (1998) reported 274 taxa representing 198 genera and 86 families in their study of Pike County Lake (220 ha). A total of 180 species collected at the Dale County Lake study area were also found to occur at Pike County Lake. This gave an index of similarity of 52.86 between these two studies. Dale County Lake and Ech Lake had 268 species in common and an index of similarity of 58.64. An index of similarity of 48.39, with 189 species in common, was calculated from the comparison of Pike County Lake and Ech Lake (Table 2). Table 2. Indicies of similarity between Dale County Lake and two study sites. Study _ Ha _ # of taxa _ Index of similarity (%) Ech Lake 162 507 58.64 with Dale County Lake 48.39 with Pike County Lake Pike County Lake 220 274 48.39 with Ech Lake 52.86 with Dale County Lake Dale County Lake 142 407 100 CONCLUSIONS After 28 months of intensive collecting, this list of 407 taxa must include nearly all species occurring at the Dale County Lake study area. Since much of the study site is a managed disturbed area, the likelihood of documenting all species of vascular plants growing at Dale County Lake is improbable. For example, sportsmen and others involved in various forms of recreation constantly alter the habitats and are likely responsible for the eradication of some species and the introduction of others. Additionally, the successfulness of the beavers in damming the creeks has caused a rapid change in wetland habitats. Fifty-three (13.0%) of the species on our checklist are non-native. Twenty-one (39.6%) of these were collected from the grassland habitat, the most disturbed habitat at the 68 Woods, et al. study area. These 21 species represent 32.3% of the total (65) collected from this habitat. In the remaining four habitats (shoreline, mesic woods, xeric woods and wetlands), which are the least disturbed, only 32 (9.4%) of the species are non-native (Table 1). According to Vitousek (1988), non-native plants constitute 6-27% of the flora from the national parks in the United States. Most of these non-native species are confined to such sites as agricultural land, roadsides, and recreation areas. This is likely true for our study area as well, since 39.6% of the non-native flora occurred in the most disturbed habitat. Basinger et al. (1997) reported the percentage of introduced species from intensively managed and non- intensively managed sites in Illinois, Missouri, and Tennessee ranged from 17.6% to 21.3% and 2.7% to 15.7%, respectively. The 13.0% of non-native species collected during this survey would suggest that the study area is non-intensively managed with a small number of introduced species. Although the study area is heavily used in various forms of recreation, the land is not managed to create wildlife habitat. Additionally, management techniques such as burning and selective logging are not practiced. With the exception of the grassland habitat, very few management practices occur. Most of the grassland habitat is located around the central office, parking lot, boat-ramp, land piers, and along the 5.6 km walking trail around the lake. Although the walking trail passes through the four least disturbed habitats, very little vegetation damage has occurred because most people remain on the trail. Since little disturbance has occurred in these habitats, the opportunity for non-native species to be introduced and become established has been minor. All three of the study sites selected to determine index of similarity are in the Eastern Red Hills Region of the Coastal Plain Providence of southeast Alabama. According to Barbour et al. (1987), two sites with an index of similarity greater than 50 represent the same community types. Based on this criterion, Ech Lake and Dale County Lake have the most similar community types (IS = 58.64). In part, this could be due the fact that these two sites are closely located to one another. Dale County Lake is 16 km northeast of Ech Lake. The Dale County Lake site is 47 km southeast of Pike County Lake and these two sites have an index of similarity of 52.86. Both of these sites consist primarily of moderately drained soils, which form a mesic habitat. The index of similarity between the Pike County Lake site, located 49 km north of Ech Lake, is 48.39. Several factors, including habitat diversity and number of species collected, likely contribute to these three sites not having higher index similarities. For example, cypress swamps and ravines, which are present at both the Dale County and Pike County sites, do not occur at Ech Lake. Additionally, most soils around Ech Lake are excessively to moderately drained resulting in extremely xeric conditions. Only 274 taxa were collected during the Pike County study. This was attributed to Hurricane Opal, which occurred during the course of the study. Extensive vegetation damage was done by the hurricane and by subsequent logging crews, which were brought in to clear potentially hazardous debris. Several normally abundant species could not be found due to the stress on the ecosystem caused by both the hurricane and the logging and burning operation conducted afterwards (Woods and Reiss 1998). It is hoped that this survey of the vascular flora of the Dale County Lake study area will provide a basis of comparison for future floristic studies at other county lakes and similar areas in Alabama. Also, the information contained in this list could be used to inform the public of the uniqueness of the plants in the area. 69 Flora of Dale County Lake ANNOTATED CHECKLIST The nomenclature, in most cases, follows that of Kartesz and Kartesz (1980). Introduced species are indicated by an asterick (*) prior to the generic name. The plant families are arranged alphabetically. Generic names and specific epithets, within each family, are also listed alphabetically. The general habitat(s) and collection number(s) are provided for each taxon. Collection numbers 8100 through 8243 represent specimens collected by the first and third authors while collection numbers 1 through 350 represent specimens collected by the second author. ACANTHACEAE Ruellia caroliniensis (J.F. Gmel.) Steud.; mesic woods; 186. ACERACEAE Acer floridanum (Chapman) Pax; mesic woods; 224. A. rubrum L.; mesic woods; 103, 305. AGAVACEAE Yucca filamentosa L.; xeric woods; 187. AIZOACEAE Mollugo verticillata L.; wetland; 8201. ALISMACEAE Sagittaria latifolia Willd.; mesic woods; 55. AMARANTFLACEAE * Alternanthera philoxeroides (Mart.) Griseb.; shoreline; 1 1. ANACARDIACEAE Rhus copallina L.; xeric woods; 225. Toxicodendron radicans (L.); mesic woods; 292. ANNONACEAE Asimina parviflora (Michx.) Dunal; mesic woods; 8150. APIACEAE *Centella asiatica (L.) Urban; grassland; 8226. Chaerophyllum tainturieri Hook.; mesic woods; 126. Ciclospermum leptophyllum (Pers.); mesic woods; 144. Cicuta mexicana Coult. & Rose; wetland; 38. Eryngium prostratum Nutt.; shoreline; 165. Hydrocotyle umbellata L.; shoreline; 8104. H. verticillata Thunb.; shoreline; 8124. Sanicula canadensis L.; wetland; 8192. S. smallii Bickn.; xeric woods; 8175. AQUIFOLIACEAE Ilex aquifolium L.; xeric woods; 8119. /. glabra (L.) Gray; mesic woods; 306. I. opaca Ait.; mesic woods; 17. I. vomitoria Ait.; mesic woods; 55. ARACEAE Arisaema triphyllum (L.); wetland; 188. 70 Woods, et al. *Colocasia esculenta (L.) Schott; wetland; 8109. Peltandra virginica (L.) Schott; shoreline; 18. ARALIACEAE Aralia spinosa L.; mesic woods; 189. ARISTOLOCHIACEAE Hexastylis arifolia (Michx.) Small; mesic woods; 127. ASCLEPIADACEAE Asclepias variegata L.; mesic woods; 167. Matelea gonocarpa (Walt.) Shinners; mesic woods; 253. ASPLENIACEAE Asplenium platyneuron (L.) Oakes ex D.C. Eat.; mesic woods; 56. Athyrium filix-femina (L.) Roth; wetland; 8125. Onoclea sensibilis L.; wetland; 294. Polystichum acrostichoides (Michx.) Schott; mesic woods; 57. Thelypteris hispidula (Decaisne) C.F. Reed; wetland; 8190. T. kunthii (Desv.) Morton; wetland; 8111. T. torresiana (Gaud.) Alston; wetland; 8195. ASTERACEAE Ambrosia artemisiifolia L.; wetland; 244. Aster paternus Cronq.; mesic woods; 39. A. pilosus Willd.; mesic woods; 58. *Baccharis halimifolia L.; mesic woods; 78. Chrysopsis mariana (L.) Ell.; xeric woods; 69. Cirsium horridulum Michx.; grassland; 145. Conyza canadensis (L.) Cronq.; mesic woods; 245. Coreopsis lanceolata L.; mesic woods; 146. C. tinctoria Nutt.; grassland; 168. Elephantopus carolinianus Raeusch.; xeric woods; 8127. E. tomentosus L.; mesic woods; 190 Erechtites hieraciifolia (L.) Raf. ex DC.; grassland; 73. Erigeron annuus (L.) Pers.; grassland; 147. E. philadelphicus L.; mesic woods; 128. E. strigosus Muhl. ex Willd.; grassland; 70. Eupatorium aromaticum L.; mesic woods; 254. E. capillifolium (Lam.) Small; mesic woods; 255. E. coelestinum L.; wetland; 79. E. fistulosum Barratt; grassland; 191 . E. rotundifolium L.; xeric woods; 8173. E. serotinum Michx.; mesic woods; 80. Euthamia tenuifolia (Pursh) Greene; mesic woods; 8 1 . Gaillardia pulchella Foug.; mesic woods; 148. Gnaphalium falcatum (Lam.) Torr. & Gray; grassland; 295. G. pensilvanicum Willd.; xeric woods edge; 8145. G. purpureum L.; mesic woods; 82. Helenium autumnale L.; wetland; 257. Helianthus angustifolius L.; mesic woods; 258. 71 Flora of Dale County Lake H. microcephalus Torr. & Gray; mesic woods; 169. H. resinosus Small; mesic woods; 71. Hieracium gronovii L.; grassland; 307. * Hypochoeris brasiliensis (Less.) Hook. & Arn.; xeric woods; 8154. Krigia caespitosa (Raf.) Chambers; grassland; 104. K. virginica (L.) Willd.; grassland; 297 . Lactuca canadensis L.; mesic woods; 72. Liatris graminifolia Michx.; xeric woods; 81 10. L. spicata (L.) Willd.; xeric woods edge; 8221. Mikania scandens (L.) Willd.; mesic woods; 19. Pityopsis aspera (Shuttlw.) Small; grassland; 193. P. graminifolia (Michx.) Nutt.; xeric woods edge; 8222. Pluchea odorata (L.) Cass.; wetland; 8207. Prenanthes serpentaria Pursh; mesic woods; 260. Pyrrhopappus carolinianus (Walt.) DC.; grassland; 129. Senecio anonymus Wood; grassland; 130. Silphium asteriscus L.; mesic woods; 40. Solidago caesia L.; mesic woods; 84. S. canadensis L.; mesic woods; 83. S. odora Ait.; mesic woods; 8231. S. petiolaris Ait.; mesic woods; 85. S. rugosa Ait.; mesic woods; 86. Soliva pterosperma (Juss.) Less.; grassland; 299. *Sonchus asper (L.) Hill; mesic woods; 226. * Taraxacum officinale Weber; grassland; 87. Vernonia angustifolia Michx.; wetland; 8116. V. gigantea (Walt.) Trel. ex Branner & Coville; mesic woods; 170. BERBERIDACEAE *Nandina domestica Thunb.; mesic woods; 105. BETULACEAE Alnus serrulata (Ait) Willd.; wetland; 19. Carpinus caroliniana Walt.; mesic woods; 194. BIGNONIACEAE Bignonia capreolata L.; mesic woods; 270. Campsis radicans (L.) Seem, ex Bureau; mesic woods ; 41. Catalpa bignonioides Walt.; mesic woods; 195. BLECHNACEAE Woodwardia areolata (L.) T. Moore; wetland; 20. BRASSICACEAE *Cardamine hirsuta L.; mesic woods; 88. C. pensylvanica Muhl. ex Willd.; mesic woods; 300. Lepidium virginicum L.; xeric woods; 42. BROMELIACEAE Tillandsia usneoides (L.) L.; mesic woods; 149. CACTACEAE Opuntia humifusa (Raf.) Raf.; xeric woods; 8106. 72 Woods, et al. CAMPANULACEAE Lobelia puberula Michx.; wetland; 74. Triodanis perfoliata (L.) Nieuwl.; mesic woods; 106. Wahlenbergia marginata (Thunb.) A. DC.; mesic woods; 107. CAPRIFOLIACEAE *Lonicera japonica Thunb.; mesic woods; 21. L. sempervirens L.; mesic woods; 301. Sambucus canadensis L.; mesic woods; 150. Viburnum dentatum L.; mesic woods; 22. V. nudum L.; mesic woods; 151. V. rufidulum Raf.; mesic woods; 8139. CARYOPHYLLACEAE *Stellaria media (L.) Vill.; mesic woods; 89. CELASTRACEAE Euonymus americanus L.; mesic woods; 131. CHENOPODIACEAE *Chenopodium ambrosioides L.; xeric woods edge; 8198. CISTACEAE Helianthemum carolinianum (Walt.) Michx.; xeric woods edge; 8163. CLUSIACEAE Hypericum hypericoides (L.) Crantz; xeric woods; 227. H. mutilum L.; shoreline; 8177. H. stans (Michx.) P. Adams & Robson; grassland; 228. Triandenum walteri (J.G. Gmel.) Gleason; shoreline; 8138. COMMEL1NACEAE *Commelina communis L.; wetland; 8118. C. virginica L.; wetland; 8211. * Murdannia keisak (Elassk.) Eland. -Maz.; wetland; 8126. CONVOLVULACEAE Cuscuta compacta Juss.; mesic woods; 261. Dichondra carolinensis Michx.; grassland; 302. Ipomoea hederifolia L.; grassland; 8212. I. pandurata (L.) G. Mey; xeric woods edge; 8147. I. purpurea (L.) Roth; xeric woods edge; 8204. Jacquemontia tamnifolia (L.) Griseb.; grassland; 8214. CORNACEAE Cornus alternifolia L. f.; mesic woods; 196. C. florida L.; mesic woods; 90. C.foemina P. Mill.; mesic woods; 8100. CUPRESSACEAE Juniperus virginiana L.; rtiesic woods; 197. CYPERACEAE Carex complanata Torr. & Hook.; mesic woods; 8156. C. debilis Michx.; mesic woods; 8137. C. frankii Kunth; wetland; 8115. C. glaucescens Ell.; wetland; 229. 73 Flora of Dale County Lake C. lupulina Willd.; wetland; 8132. C. lurida Wahlenb.; wetland; 44. C. vulpinoidea Michx.; wetland; 316. Cyperus erythrorhizos Muhl.; shoreline; 8174. C. globulosus Aubl.; mesic woods; 198. C. pseudovegetus Steud.; mesic woods; 171. C. sesquiflorus (Torr.) Mattf. & Kukenth.; shoreline; 8103. C. strigosus L.; mesic woods; 199. C. virens Michx.; wetland; 8168. Eleocharis microcarpa Torr.; shoreline; 8146. E. obtusa (Willd.) Schultes; shoreline; 8123. E. tuberculosa { Michx.) Roemer & Schultes; mesic woods; 132. Fimbristylis autumnalis (L.) Roemer & Schultes; shoreline; 8185. F. dichotoma (L.) Vahl; shoreline; 8184. Rhynchospora cephalantha Gray; mesic woods; 340. R. mixta Britt, ex Small; mesic woods; 91, 8129. Scirpus cyperinus (L.) Kunth; wetland; 45. Scleria triglomerata Michx.; xeric woods edge; 8157. DIOSCOREACEAE Dioscorea quaternata (Walt.) J.F. Gmel.; shoreline; 8203. EBENACEAE Diospyros virginiana L.; mesic woods; 262. ELAEAGNACEAE * Elaeagnus pungens Thunb.; mesic woods; 200. ERICACEAE Kalmia latifolia L.; mesic woods; 172. Lyonia lucida (Lam.) K. Koch; mesic woods; 46. Oxydendrum arboreum (L.) DC.; mesic woods; 173. Rhododendron canescens (Michx.) Sweet; mesic woods; 108. Vaccinium arboreum Marsh.; mesic woods; 133. V. corymbosum L.; mesic woods; 23. V. elliottii Chapman; mesic woods; 134. V. stamineum L.; mesic woods; 135. EUPHORBIACEAE Acalypha gracilens Gray; xeric woods edge; 8161. Chamaesyce maculata (L.) Small; grassland; 8236. Croton glandulosus L.; xeric woods edge; 8153. Euphorbia corollata L.; xeric woods; 24. * Phyllanthus uninaria L.; grassland; 8233. Poinsettia heterophylla (L.) Klotzsch & Garzke; xeric woods edge; 68. *Sapium sebiferum (L.) Roxb.; mesic woods; 201. Sebastiania fruticosa (Bartr.) Fern.; mesic woods; 174. Tragia smallii Shinners; xeric woods edge; 8152. T. urticifolia Michx.; xeric woods edge; 8160. FABACEAE * Albizia julibrissin Durz.; mesic woods; 47. 74 Woods, et al. Amorpha fruticosa L.; shoreline; 8169. Apios americana Medic.; wetland; 230. Cassia fasciculata Michx.; xeric woods; 246. C. nictitans L.; mesic woods edge; 8235. Cercis canadensis L.; mesic woods; 304. Clitoria mariana L.; grassland; 152. Crotalaria rotundifolia (Walt.) Poir.; wetland; 313. Desmodium laevigatum (Nutt.) DC.; xeric woods edge; 8242. D. lineatum DC.; mesic woods; 8234. D. obtusum (Muhl. ex Willd.) DC.; mesic woods edge; 8241. D. strictum (Pursh) DC.; xeric woods; 62. *Erythrina herbacea L.; mesic woods; 153. Galactia volubilis (L.) Britt.; mesic woods; 23 1. * Kummerowia striata (Thunb.) Schindl.; grassland; 8238. *Lespedeza cuneata (Dum.-Cours.) G. Don; grassland; 48. L. procumbens Michx.; mesic woods; 109. L. stuevei Nutt.; xeric woods edge; 8240. L. virginica (L.) Britt.; xeric woods edge; 8239. *Medicago polymorpha L.; grassland; 272. *Melilotus alba Medic.; grassland; 154. *Pueraria lobata (Willd.) Ohwi; mesic woods; 232. Schrankia microphylla (Dry.) J.F. Macbr.; mesic woods; 155. Strophostyles umbellata (Muhl. ex Willd.) Britt.; mesic woods; 248. Stylosanthes biflora (L.) B.S.P.; xeric woods edge; 8187. Tephrosia spicata (Walt.) Torr. & Gray; grassland; 233. * Trifolium campestre Schreb.; grassland; 92. *T. dubium Sibthorp; grassland; 273. *T. repens L.; grassland; 274. *T. vesiculosum Savi; grassland; 49. * Vida saliva L.; mesic woods; 1 10. FAGACEAE Fagus grandifolia Ehrh.; mesic woods; 202. Quercus alba L.; mesic woods; 203. Q. falcata Michx.; mesic woods; 320. Q. hemisphaerica Bartr.; mesic woods; 204. Q. nigra L.; mesic woods; 206. Q. shumardii Buckl.; mesic woods; 8120. Q. stellata Wang.; mesic woods; 264. GENTIANACEAE Gentiana catesbaei Walt.; mesic woods; 50. GERANIACEAE Geranium carolinianum L.; mesic woods; 111. HAMAMELIDACEAE Hamamelis virginiana L.; mesic woods; 8140. Liquidambar styraciflua L.; xeric woods; 207. HIPPOCASTANACEAE 75 Flora of Dale County Lake Aesculus pavia L.; mesic woods; 1 12. HYDROPHYLLACEAE Hydrolea quadrivalvis Walt.; wetland; 235. IRIDACEAE Sisyrinchium angustifolium P. Mill.; wetland; 8216. *S. exile Bickn.; grassland; 8102. S. rosulatum Bickn.; mesic woods; 156. JUGLANDACEAE *Carya illinoensis (Wang.) K. Koch; wetland; 25. C. pallida (Ashe) Engl. & Graebn.; xeric woods; 236. C. tomentosa (Poir.) Nutt.; mesic woods; 208. JUNCACEAE Juncus acuminatus Michx.; shoreline; 8159. J. effusus L.; wetland; 136. J. marginatus Rostk.; wetland; 26. J. polycephalus Michx.; wetland; 8128. J. repens Michx.; shoreline; 8112. Luzula acuminata Raf.; mesic woods; 8220. LAMIACEAE *Lamium amplexicaule L.; mesic woods; 93. Lycopus virginicus L.; wetland; 27, 8191. Monarda punctata L.; xeric woods edge; 8227. Pycnanthemum incanum (L.) Michx.; mesic woods; 237. Salvia lyrata L.; mesic woods; 157. Scutellaria elliptica Muhl.; mesic woods; 158. S. incana Biehler; wetland; 8206. S. integrifolia L.; wetlands; 51. Stachys floridana Shuttlw.; mesic woods; 28. LAURACEAE Persea borbonia (L.) Spreng.; mesic woods; 321. Sassafras albidum (Nutt.) Nees; mesic woods; 209, 276. LENTIBULAR1ACEAE Utricularia biflora Lam.; aquatic; 8108. LILIACEAE Nothoscordum bivalve (L.) Britt.; grassland; 113. LOGANIACEAE Gelsemium sempervirens (L.) St. Hil.; mesic woods; 94. Mitreola petiolata (Gmel.) Torr. & Gray; wetland; 8224. Polypremum procumbens L.; grassland; 52. Spigelia marilandica L.; mesic woods; 159. LYTHRACEAE Rotala ramosior (L.) Koehne; wetland; 8209. MAGNOLIACEAE Liriodendron tulipifera L.; mesic woods; 137. Magnolia acuminata (L.) L.; mesic woods; 8143. 76 Woods, et al. M. grandiflora L.; wetland; 53. M. virginiana L.; mesic woods; 160. MALVACEAE Hibiscus aculeatus Walt.; wetland; 250. *Sida rhombifolia L.; xeric woods; 8196. MELASTOMATACEAE Rhexia mariana L.; wetland; 8219. R. virginica L.; mesic woods; 54. MELIACEAE *Melia azedarach L.; mesic woods; 238. MORACEAE *Morus alba L.; mesic woods; 212. M. rubra L.; mesic woods; 213. MYRJCACEAE Myrica cerifera L.; wetland; 29. NYSSACEAE Nyssa sylvatica Marsh.; mesic woods; 214. OLEACEAE Chionanthus virginicus L.; mesic woods; 8114. Fraxinus caroliniana P. Mill.; mesic woods; 265. F. pennsylvanica Marsh.; wetland; 8170. * Ligustrum japonicum Thunb.; wetland; 8200. *L. sinense Lour.; mesic woods; 12. Osmanthus americanus (L.) Benth. & Hook. f. ex Gray; mesic woods; 216. ONAGRACEAE Ludwigia alternifolia L.; mesic woods; 30. L. bonariensis (M. Micheli) Hara.; mesic woods; 3 1. L. decurrens Walt.; shoreline; 8214. L. repens Forst.; wetland; 8189. L. spathulata Torr. & Gray; shoreline; 8232. Oenothera biennis L.; grassland; 277. O. laciniata Hill; grassland; 95. *0. speciosa Nutt.; grassland; 8213. ORCHIDACEAE Tipularia discolor (Pursh) Nutt.; mesic woods; 60. OROBANCHACEAE Conopholis americana (L.) Wallr.; mesic woods; 278. OSMUNDACEAE Osmunda cinnamomea L.; mesic woods; 138. O. regalis L.; mesic woods; 217. OXALIDACEAE Oxalis dillenii Jacq.; mesic woods; 1 14. PASSIFLORACEAE Passiflora incarnata L.; grassland; 161. P. lutea L.; mesic woods; 313. PHYTOLACCACEAE 77 Flora of Dale County Lake Phytolacca americana L.; mesic woods; 162. PINACEAE Pinus glabra Walt.; mesic woods; 3 12. P. taeda L.; grassland 96. PLANTAGINACEAE Plantago aristata Michx.; xeric woods edge; 8148. *P. lanceolata L.; grassland; 280. P. virginica L.; mesic woods; 1 15. PLATANACEAE Platanus occidentalis L.; mesic woods; 218. POACEAE Agrostis hiemalis (Walt.) B.S.P.; grassland; 283. Andropogon ternarius Michx.; grassland; 28. A. virginicus L.; grassland; 282. Arundinaria gigantea (Walt.) Muhl.; wetland; 284. Axonopus affinis Chase; grassland; 285. *Briza minor L.; grassland; 286. Bromus catharticus Vahl; mesic woods; 8172. Cenchrus echinatus L.; grassland; 219. Chasmanthium sessiliflorum (Poir.) Yates; mesic woods; 75. Cynodon dactylon (L.) Pers.; grassland; 8176. Dactyloctenium aegyptium (L.) Beauv.; grassland; 8230. Dichanthelium commutatum (Schultes) Gould; mesic woods; 33. D. dichotomum (L.) Gould; shoreline; 8136. D. scoparium (Lam.) Gould; mesic woods; 32, 8166. *Digitaria ciliaris (Retz.) Koel.; grassland; 8180. Eragrostis spectabilis (Pursh) Steud.; xeric woods edge; 8155. Erianthus contortus Baldw.; grassland; 1 16. Festuca arundinacea Schreb.; grassland; 1 17. Gymnopogon ambiguus (Michx.) B.S.P.; wetland; 8229. Leersia oryzoides (L.) Sw.; shoreline; 8130. L. virginica Willd.; shoreline; 8197. Panicum anceps Michx.; shoreline; 8237. P. dichotomiflorum Michx.; mesic woods; 315. Paspalum distichum L.; shoreline; 8243. P. laeve Michx.; grassland; 34. *P. notatum Fluegge; grassland; 35. P. setaceum Michx.; xeric woods edge; 8158. *P. urvillei Steud.; grassland; 36. *Poa annua L.; grassland; 266. Sacciolepis striata (L.) Nash; shoreline; 8181. Sphenopholis obtusata (Michx.) Scribn.; xeric woods; 8117. Sporobolus clandestinus (Biehler) A.S. Hitchc.; xeric woods; 8131. *S. indicus (L.) R. Br.; mesic woods; 8228. Tridens Jlavus (L.) A.S. Hitchc.; grassland; 1 18. POLEMONIACEAE 78 Woods, et al. Phlox Carolina L.; mesic woods; 139. POLYGALACEAE Polygala grandijlora Walt.; mesic woods; 163. POLYGONACEAE Polygonum densiflorum Meisn.; wetland; 8202, 8225. Rumex crispus L.; grassland; 311. R. hastatulus Baldw. ex Ell; grassland; 1 19. POTAMOGETONACEAE Potamogeton diversifolius Raf.; aquatic; 8210. PTER1DACEAE Pteridium aquilinum (L.) Kuhn; mesic woods; 8107. RANUNCULACEAE Clematis reticulata Walt.; wetland; 37. C. virginiana L.; mesic woods; 25 1 . Xanthorhiza simplicissima Marsh.; mesic woods; 287. RHAMNACEAE Berchemia scandens (Hill) K. Koch; mesic woods; 8101. Ceanothus americanus L.; mesic woods; 176. ROSACEAE Alchemilla microcarpa Boiss. & Reut.; grassland; 317. Amelanchier arborea (Michx. f.) Fern.; mesic woods; 289. Aronia arbutifolia (L.) Pers.; mesic woods; 122, 339. Crataegus j, lava Ait.; xeric woods; 8144. C. marshallii Egglest.; mesic woods; 140. C. uniflora Muenchh.; xeric woods; 8167. *Duchesnea indica (Andr.) Focke; mesic woods; 177. Prunus angustifolia Marsh.; mesic woods; 242. P. caroliniana (P. Mill.) Ait.; mesic woods; 97. P. serotina Ehrh.; mesic woods; 120. P. umbellata Ell.; xeric woods; 8199. *Pyrus communis L.; mesic woods; 267 . Rosa Carolina L.; mesic woods; 178. Rubus betulifolius Small; wetland; 8165. R. cuneifolius Pursh; mesic woods; 121. R. flagellaris Willd.; xeric woods edge; 8135. RUBIACEAE Cephalanthus occidentalis L.; wetland; 179. Diodia teres Walt.; grassland; 8162. D. virginiana L.; grassland; 8164. Galium aparine L.; mesic woods; 268. G. circaezans Michx.; xeric woods edge; 8151. G. pilosum Ait.; grassland; 180. Hedyotis crassifolia Raf.; grassland; 98. H. procumbens (Walt, ex J.F. Gmel.) Fosberg; mesic woods; 309. H. purpurea (L.) Torr. & Gray; mesic woods; 3 1 0. Mitchella repens L.; mesic woods; 76. 79 Flora of Dale County Lake * Richardia scabra L.; mesic woods; 99. SALICACEAE Salix nigra Marsh.; wetland; 8105. SAXIFRAGACEAE Decumaria barbara L.; mesic woods; 13. Hydrangea quercifolia Bartr.; mesic woods; 290. Itea virginica L.; shoreline; 14. Penthorum sedoides L.; wetland; 8182. SCHIZAEACEAE *Lygodium japonicum (Thunb.) Sw.; mesic woods; 15. SCROPHULAR1ACEAE Agalinis fasciculata (Ell.) Raf.; mesic woods; 77. Aureolaria j lava (L.) Farw.; xeric woods edge; 8141. A. virginica (L.) Pennell; mesic woods; 181. Linaria canadensis (L.) Dum.-Cours.; grassland; 123. Linder nia dub ia (L.) Pennell; shoreline; 8178. *Mazus pumilus (Burm. f.) Steenis; mesic woods; 291. Micranthemum umbrosum (Walt.) Blake; wetland; 8188. * Veronica arvensis L.; mesic woods; 269. SMILACACEAE Smilax bona-nox L.; mesic woods; 252. S. glauca Walt.; mesic woods; 243. S. lasioneuron Hook.; xeric woods; 8142 S. laurifolia L. ; wetlands; 308. S. pumila Walt.; mesic woods; 61. S. rotundifolia L.; xeric woods; 220. S. smallii Morong; mesic woods; 182. SOLANACEAE Solanum americanum P. Mill.; mesic woods; 16. S. carolinense L.; mesic woods; 222. STYRACACEAE Styrax americana Lam.; mesic woods; 141. S. grandifolia Ait.; mesic woods; 142. ULMACEAE Celtis tenuifolia Nutt.; xeric woods; 8113. Ulmus alata Michx.; xeric woods; 8133. URT1CACEAE Boehmeria cylindrica (L.) Sw.; wetland; 223. VALERIANACEAE Valerianella radiata (L.) Dufr.; mesic woods; 124. VERBENACEAE Callicarpa americana L.; mesic woods; 183. * Verbena brasiliensis Veil.; grassland; 143. *V. rigida Spreng.; grassland; 184. VIOLACEAE Viola affinis Le Conte; mesic woods; 100. 80 Woods, et al. V. palmata L.; mesic woods; 101. V. primulifolia L.; mesic woods; 125. V. rafinesquii Greene; mesic woods; 102. VITACEAE Parthenocissus quinquefolia (L.) Planch.; mesic woods; 185. Vitis aestivalis Michx.; xeric woods edge; 8149. V. cinerea Engelm. ex Millard; xeric woods; 8122. V. rotundifolia Michx.; mesic woods; 164. XYR1DACEAE Xyris difformis Chapman; wetland; 8208. X. jupicai L.C. Rich.; shoreline; 8183. ACKNOWLEDGEMENTS The authors would like to thank The Alabama Department of Conservation and the Ozark Department of Parks and Recreation for granting permission to conduct this study at Dale County Lake. Also, we are greatful to Hannelore Rundell, Tiffany Pennington and Brian Martin for their help during this project. The second author would like to thank the College of Arts and Sciences at Troy State University for awarding him a Chancellor's Fellowship. Support for this project was provided by The Alabama Department of Public Health ALERT Grant. LITERATURE CITED Barbour, M.G., J.H. Burke, and W.D. Pitts. 1987. Terrestrial plant ecology, 2nd ed. The Benjamin/Cummings Company, Menlo Park, California. Basinger, M.A., J.S. Huston, R.J. Gates, and P.A. Robertson. 1997. Vascular Flora of Horseshoe Lake Conservation Area, Alexander County, Illinois. Castanea 62(2): 82- 99. Brown, M. 1999. Weather data for the years 1997 and 1998. Personal communication with the National Weather Service. Clewell, A.F. 1985. Guide to the vascular plants of the Florida Panhandle. Florida State University Press, Tallahassee. Cronquist, A. 1980. Vascular flora of the southeastern United States. Volume I. Asteraceae. The University of North Carolina Press, Chapel Hill. Crouch, V.E. and M.S. Golden. 1997. Floristics of a bottomland forest and adjacent uplands near the Tombigbee River, Choctaw County, Alabama. Castanea 62(4): 219-238. Diamond, A.R and J.D. Freeman. 1993. A checklist of the vascular flora of Conecuh County, Alabama. Sida 15(4):623-638. Godfrey, R.K. 1988. Trees, shrubs and woody vines of Northeastern Florida and adjacent Georgia and Alabama. University of Georgia Press, Athens. Godfrey, R.K. and J.W. Wooten. 1979. Aquatic and wetland plants of the southeastern United States. Monocotyledons. University of Georgia Press, Athens. Godfrey, R.K. and J.W. Wooten. 1981. Aquatic and wetland plants of the southeastern United States. Dicotyledons. University of Georgia Press, Athens. Harper, R.M. 1943. Forests of Alabama. Alabama Geol. Surv. Monograph 10: 1-230. 81 Flora of Dale County Lake Hitchock, A.S. 1971. Manual of the grasses of the United States. Volumes I & II. General Publishing Company, Ltd., Tornonto. Isely, A. 1990. Vascular flora of the southeastern United States. Volume III, Part 2. Leguminosae (Fabaceae). The University of North Carolina Press, Chapel Hill. Kartesz, J.T. and R. Kartesz. .1980. A synonymized checklist of the vascular flora of the United States, Canada and Greenland. The University of North Carolina Press, Chapel Hill. Mohr, C. 1901. Plant life of Alabama. Government Printing Office. Washington D.C. Mount, R.M. and A.R. Diamond. 1992. Final survey of the fauna and flora of Fort Rucker, Alabama. Auburn University. Mueller-Dombois, D. and H. Ellenbery. 1974. Aims and methods of vegetation ecology. Wiley, New York. Radford, A.E., H.E. Aides and C.R. Bell. 1968. Manual of the vascular Bora of the Carolinas. The University of North Carolina Press, Chapel Hill. Rundell, H. and A.R. Diamond. 1999. Noteworthly collections from Alabama. Castanea 64(4):355-356. Rundell, H. and M. Woods. 2001. The vascular flora of Ech Lake, Alabama. Castanea 66(4): in press. Small, J.K. 1933. Manual of the southeastern flora. The University of North Carolina Press, Chapel Hill. Vitousek, P.M. 1988. Diversity and biological invasions of oceanic islands. P. 181-189 In: Wilson, E.O. and F.M. Peters (eds.). Biodiversity, Chapter 13, National Academic Press, Washington, D.C. Woods, M. and J. Reiss. 1998. The vascular flora of Pike County Lake, Alabama. J. Alabama Acad. Sci. 69(3):300-3 14. 82 NOTES NOTES INSTRUCTIONS TO AUTHORS Editorial Policy: Publication of the Journal of the Alabama Academy of Science is restricted to members. Membership application forms can be obtained from Dr. A. Priscilla Holland. Office of Research, UNA Box 5121, University of North Alabama, Florence, AL 35632-0001. 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