.2 . .i V , +4533... 234$ 1.. .x 1.. I {:1 3.19. . - u .945... u" 5.9., WA“, 96... i? . .r‘. .n .. (bl! [Law Li. .3, .. it .t) ‘80 it “NY . it'll Hahn. ‘ a. .‘I‘I‘ \ Iii}... I 1:.7 ‘ . .3 .1 .32... ..-&a$i§§$§3§% “fiafifipé Iuniniljiifl'lit‘iTimmy:m 9 01770 LIBRARY E Michigan State I Universlty | This is to certify that the dissertation entitled Transformation Studies of Creeping Bentgrass (Agrostis palustris Huds.) Using an Elm Chitinase Gene for Disease Resistance presented by Benli Chai has been accepted towards fulfillment of the requirements for d Ph.D- degree in Wes Major professor Date g” ’3‘ " O\O\ MS U is an Affirmative Action/Equal Opportunity Institution 0-1277 1 REMOTE STORAGE ES F: PLACE IN RETURN BOX to remove this checkout from your record. TO AVOID FINES return on or before date due. DATE DUE DATE DUE DATE DUE n363y1 20:: Blue 10/13 pJCIRC/DateDueForrns_2013.nndd - p95 TRANSFORMATION STUDIES OF CREEPING BENTGRASS (AGROSTIS PALUSTRIS HUDS.) USING AN ELM CHITINASE GENE FOR DISEASE RESISTANCE BY Benli Chai A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Crop and Soil Sciences 1999 ABSTRACT TRANSFORMATION STUDIES OF CREEPING BENTGRASS (AGROSTIS PALUSTRIS HUDS.) USING AN ELM CHITINASE GENE FOR DISEASE RESISTANCE BY Benli Chai Creeping bentgrass (Agrostis palustris Huds.) is a major cool season turfgrass species used for golf greens,. tees, and fairways. This important turf species is susceptible to a variety of fungal diseases including brown patch (Rhizoctonia spp.). Genetic transformation technology has made it possible to introduce cloned genes into plant genomes for the purpose of conferring disease resistance and other economically important traits. Among the available disease resistance genes, plant chitinase genes have proven effective in delivering fungal resistance in a number of transgenic plants. An effective microprojectile- mediated transformation system was used in our studies to produce transgenic creeping bentgrass (Agrostis palustris Huds.) plants transformed with a Elm Chitinase (H82) gene. The selectable marker gene bar (phosphinothricin acetyl- transferase, PAT) and the nonselectable H82 gene on separate plasmids were co-introduced into the creeping bentgrass genome, when embryogenic calli of creeping bentgrass were bombarded with tungsten particles coated with DNA of both plasmids. Transgenic plants carrying the H82 gene were identified from bar gene-transformed plants, which were resistant to the herbicides bialaphos and Igniten“. A total of eleven H82 transformed lines were obtained from forty-six independent lines transformed with the bar gene. The frequency of unlinked cotransformation of bar and the H82 was 23.9%, which was comparable to that of other studies. Expressions of the H82 gene was detected I from some transgenic lines of creeping bentgrass at both the transcriptional and the translational levels. Two of the H82 expressing lines exhibited enhanced resistance to Rhizoctonia solani. DEDICATION Dedicated to my dear wife Hong Zhang. If it were not for her love, support, motivation, and personal sacrifices, this dissertation may have never been completed. To my daughters, Yan and Chelsea, who are my inspirations. iv ACKNOWLEDGEMENTS I would like to acknowledge all those who helped me to make this dissertation a reality. My appreciation and gratitude go to Dr. Mariam Sticklen, my major professor, for her encouragement, patience, and valuable advice. I am very thankful to other members of my guidance committee: Dr. Joseph Vargas, Dr. Joseph Saunders, and Dr. Richard Ward for the valuable advice throughout my Ph.D. program. My thanks go to the United States Golf Association for providing me part of the financial support. Special thanks to Dr. Heng Zhong, Dr. Chien—An Liu, and Mr. Don Warkentin for being my mentors and colleagues. TABLE OF CONTENTS LIST OF TABLES. LIST OF FIGURES. CHAPTER 1 TURFGRASS IN VITRO CULTURE AND GENETIC ENGINEERING Abstract. Introduction. . Turfgrass in vitro Culture. Genetic Transformation. Future Opportunities in Turfgrass Biotechnology. CHAPTER 2 CHITINASE GENES AND PLANT DISEASE RESISTANCE Introduction. Chitinase in Plants. . . Engineering Fungal Resistance Using Chitinase Genes. CHAPTER 3 GENETIC TRANSFORMATION OF CREEPING BENTGRASS (AGROSTIS PALUSTRIS HUDS.) USING A DUTCH ELM CHITINASE GENE Abstract. Introduction. Material and Methods. Results. Discussion. vi vii viii (13wa 59 .60 62 68 7O 72 .89 .108 LIST OF TABLES Table 1. Turfgrass regeneration from in vitro culture .16 Table 2. Bialaphos Selection and Plant Regeneration after Bombardment .90 Table 3. Gene integration and expression in .99 bialaphos resistant transgenic lines. vfi Figure Figure Figure Figure Figure Figure Figure Figure Figure Figure Figure 10. 11. LIST OF FIGURES Schematic diagrams of the plasmids, pJSlOl and pKYLX7l-H82. PCR detecting bar gene in creeping bentgrass transgenic plants. PCR detecting H82 gene in creeping bentgrass transgenic plants. PCR detecting mtlD gene in creeping bentgrass transgenic plants. . DNA Slot blot analyses Southern blot analyses of transgenic creeping bentgrass for mtlD gene insertions. Southern blot analyses of transgenic creeping bentgrass for H82 gene insertions. Correlation between the copy numbers of gene insertion. RT-PCR detecting H82 gene transcripts Northern blot analyses of creeping bentgrass transgenic plants. Western blot analyses of the transgenic creeping bentgrass expressing H82 gene. viii 73 .93 94 .95 .96 .100 .101 102 105 106 .107 CHAPTER ONE TURFGRASS IN VITRO CULTURE AND GENETIC ENGINEERING (LITERATURE REVIEW) ABSTRACT During the past two decades, substantial progress has been made in applying modern biotechnology in turfgrass genetic improvement. Applications of biotechnology have assisted turfgrass breeding programs in several different ways. This dissertation chapter reviews the following advances made in turfgrass biotechnology (i) in vitro manipulations for regenerable tissue culture, (11) genetic engineering by DNA transfer techniques. Further achievements in turfgrass biotechnology hold promise for solving problems that challenge the turfgrass industry. INTRODUCTION Turfgrass includes grass species that are suitable to be used as turf, i.e., a covering of vegetation, plus the matted, upper stratum of earth filled with roots and rhizomes (Beard, 1973). About 40 grass species are used as turf because of their ability to persist under regular mowing (Duble, 1996). The characteristics of low growth habit, prostate and creeping tendency, high shoot density and medium fine leaf textures have been used in selecting grasses for turf. While contributing to soil development, stabilization and improvement, as well as erosion control, turfgrass plants are also used to beautify the earth, enrich our lives, and provide recreation and enjoyment for people. Turfgrass is maintained on lawns, estates, parks, golf courses, playing fields and public grounds, which cover about 14 million hectares of the land in the United States (Duble, 1996). The turfgrass industry also plays an important role in the U.S. economy with over $580 million in annual seed sales, second only to hybrid seed corn (Zea mays L.) (Kidd, 1993; Lee, 1996). Since the establishment of turfgrass culture in North America and the rest of the world, cultivar improvement has always been a central issue. As the result of the persistent efforts of breeders, significant achievements have been made in turfgrass breeding. Over 245 warm-season and cool-season cultivars have been registered since 1946 in the U.8.A. (Lee, 1996). These achievements have contributed to the establishment and growth of the turfgrass industry and the development of turfgrass science (Beard, 1973). As in most other crops, turfgrass improvement traditionally has relied on conventional breeding methods, in which the accessible genetic material is restricted by sexual reproduction. This situation has raised the necessity of exploring other approaches for more efficient achievements. Modern plant biotechnology is based on our current understanding of biological systems at both the cellular and molecular levels and has become increasingly associated with genetic engineering and other in vitro cellular manipulations of plants (Krans, 1989). Biotechnology is being used to supplement and complement traditional methods of plant improvement by bridging the individual gene pools in nature (Vasil, 1995) and thereby creating opportunities for breeders to utilize the genetic material from heterologous sources. Introgression of genetic traits from unrelated plant species or even other kingdoms has become a realistic breeding objective (Asay and Sleper, 1989; DeBlock, 1993). TURFGRASS IN VITRO CULTURE Plant regeneration systems are critical in turfgrass biotechnology. An efficient in vitro shoot or embryo regeneration is a necessary step to recover genetically altered material, somaclonal variation, haploids, and existing germplasm with unique features, as well as for micropropagation and aseptic storage of valuable germplasm. Cell and Protoplast Cultures A common difficulty with in vitro culture of turfgrasses and other grass species is the lack of natural ability for secondary growth by cambium or cambium—like tissues in mature and differentiated explants. Slow progress in early programs was largely due to the improper use of mature and differentiated tissues as the explant material (Vasil and Vasil, 1994). kéy strategies for establishing regenerable cell cultures for grass species were reviewed and generalized by Vasil (1995) as follows: (1). choose explants made of meristematic tissues and undifferentiated cells such as immature embryos or seeds, leaf base meristems, and meristematic segments of young inflorescences; (2). use culture medium supplemented with high concentrations of strong auxins, such as 2,4-dichlor0phenoxyacetic acid (2,4—D) and 3,6-dichloro-o-anisic acid (dicamba) for inducing embryogenic calli; (3). use embryogenic calli-derived cell suspensions for protoplast isolation. These strategies have led to the development of successful regeneration systems for all major turfgrass species. A pioneer study in plant regeneration of grass species was the establishment of an embryogenic culture of Italian ryegrass (Lolium multiflorum Lam.) from immature embryos (Dale, 1980). This study and that of rice and sorghum [Sorghum bicolor (L.) Moench] provided the first observations of somatic embryogenesis in grass species (Vasil and Vasil, 1994). Efforts have been made to establish regenerable cultures from diverse turfgrass explant material (Table 1).‘ Although direct somatic embryogenesis was achieved in orchardgrass (Dactylis glomerata L.) from mesophyll cells (Hanning and Conger, 1982; Conger et al., 1983; Trigiano et al., 1989), plant regeneration from embryogenic callus is the single most important path for turfgrass, as many major turfgrass species have been regenerated in this way. The culture medium requirements for turfgrasses are similar to that for other grass species. Generally high salt nutrient solutions such as the Murashige and Skoog (M8) medium (Murashige and Skoog, 1962), the most frequently used medium for turfgrass, and modified CC-medium (Potrykus et al., 1979; Nielsen and Knudsen, 1993) with 3 to 12% sucrose and high concentrations of strong auxins such as 2,4—D and dicamba have been used. Cytokinins, such as 6-benzylaminopurine (6- BA) and N—(2-furanylmethyl)—1 H-purin—6-amine (kinetin), at low concentrations, in combination with auxins were often used in turfgrass species to promote callus initiation. Addition of casein hydrolysate to the culture medium was found to be beneficial to embryogenic callus initiation (Artunduaga et al., 1988). Proline and glutamine have stimulatory effects on callus induction among the organic N compounds tested (Shetty and Asano, 1991). Embryogenic turfgrass callus is characteristically friable, somewhat organized, and generally white to light yellow in color. Because of a low frequency of occurrence (Vasil and Vasil, 1994) and the heterogeneous nature of embryogenic cultures, selection during subculture has been effective in enrichingl embryogenic cultures (Krishnaraj and Vasil, 1995). Embryogenic cell suspensions provide the only source of totipotent cells to isolate protoplasts from grass species (Vasil, 1988; Potrykus, 1990), in addition to being the direct targets for genetic transformation. Regenerable cell suspensions are established by selectively transferring embryogenic calli to liquid medium. Finely dispersed and fast-growing cell suspensions are maintained in liquid media with high levels of auxins and subcultured every 3 to 7 d. Most turfgrass embryos only develop to either a globular or early scutellar stage in the liquid medium, as is also the case for most other gramineous species (Vasil, 1985, 1987). Cell suspensions need to be plated onto semi—solid media for further embryo development and plant regeneration. The only exception was found in orchardgrass, where embryos developped to reach a germinable stage in a single regime of liquid medium (Gray et al., 1984; Gray and Conger, 1985; Conger et al., 1989). Successful establishment of embryogenic cell suspensions and subsequent plant regeneration have been reported in several major turfgrass species, including creeping bentgrass (Terakawa et al., 1992; Hartman et al., 1994; Lee et al., 1995), tall fescue (Festuca arundinacea Schreber) (Rajoelina et al., 1990; Takamizo et al., 1990; Dalton 1988a, b, 1993; Wang et al. 1992, 1995; Ha et al., 1992; Dalton et al., 1995), red fescue (Spangenberg et al., 1994; Wang et al., 1995), meadow fescue (Wang et al., 1993a), redtop grass (Agrostis alba L.) (Asano and Sugiura, 1990); Kentucky bluegrass (Nielsen et al., 1993; Nielsen and Knudsen, 1993), Italian ryegrass (Rajoelina et al., 1990; Dalton, 1988b, 1993; Wang et al., 1993b, 1995), perennial ryegrass (Dalton, 1988a, b; Zaghmout and Torello, 1992; Wang et al., 1993b, 1995; Olesen et al., 1995a), and the hybrid ryegrass (L. x boucheanum) (Wang et al., 1993b) (Table 1.). In most of these studies, embryogenic cell suspensions were used for protoplast isolation and subsequent plant regeneration. One major problem in cell suspension cultures is the rapid decline of regenerability over time. To retain high levels of green—plant regeneration frequency, cryopreservation of suspension cultures in liquid N has been successfully introduced and optimized for perennial ryegrass, Italian ryegrass, tall fescue, and red fescue (Wang et al., 1993b; Spangenberg et al., 1994; Wang et al., 1995). Protoplasts are useful for multiple manipulations in turfgrass biotechnology. Plants have been recovered from protoplasts of several turfgrass species (Table 1.). Although having many advantages for in vitro manipulations, protoplasts are still considered the most difficult tissue- culture explants from which to recover plantlets. Embryogenic callus-derived, well-dispersed, finely grained, and fast- growing cell suspensions are required as the donor material to isolate competent protoplasts. The age of the donor-cell suspensions is very critical to the competency of protoplasts (Takamizo et al., 1990; Nielsen et al., 1993). To retain morphogenic potential of donor suspensions, addition of amino acids, such as L—proline, and an osmoticum, such as sorbitol, was beneficial (Asano and Sugiura, 1990). Using high quality enzymes for cell-wall hydrolyzation, improves protoplast isolation. In addition to the basic requirements for protoplast culture in terms of the need for nutrients, growth regulators, and osmotic pressure, the medium volume per cell ratio (plating density) and other environmental factors need to be adjusted for specific genotypes. Agarose embedding of protoplasts (bead-type culture) was used to promote plant regeneration from protoplasts (Asano and Sugiura, 1990; Takamizo et al., 1990; Wang et al., 1992, 1995). Some special treatments have been used in turfgrass species for a better regeneration response. The feeder layer system, in which protoplasts are cocultured with a layer of physically separated “nurse” or “feeder cells”, has been found effective for increasing plating efficiency of protoplasts and is necessary for the formation of protoplast-derived callus and plant regeneration in creeping bentgrass (Lee et al., 1995), tall fescue, red fescue and meadow fescue (Wang et al., 1995), and Italian ryegrass, perennial ryegrass, and L. x bouceanum (Wang et al., 1995; Olesen et al., 1995a). This method was frequently used for other turfgrass species (Takamizo et al., 1990; Ha et al., 1992; Wang et al., 1992). Proline in proliferation medium enhanced regeneration of Kentucky bluegrass from protoplasts (Takamizo et al., 1990). Despite the remarkable success that has been achieved in turfgrass species, protoplast culture still remains difficult in practice because of the intensive labor involved, significant genotype-dependence in culture establishment, and relatively short longevity of the donor-cell suspensions, problems of albinism, low frequency in plant regeneration from protoplasts, and difficulties in selection after genetic manipulations. Manipulations of culture conditions provide strong potential to overcome the recalcitrance caused by epigenetic effects, as indicated by Halperin (1986). Shoot Apex Culture The shoot apex is a small, distalmost portion of the shoot, typically including such regions as the apical meristem, three to six lateral primordia, and a subapical region (Medford, 1992). Studies of some grass species have found that the development of their shoot apices can be reprogrammed by in vitro culture. This developmental plasticity was even obvious in the early studies when fertile plants were recovered directly from shoot apices of maize (Raman et al., 1980), Italian ryegrass, perennial ryegrass, meadow fescue, tall fescue, and red fescue (Dale, 1975, 1977a, b). The more recent studies found that in vitro culture regimes could affect shoot apex development in more profound ways. Somatic embryos can be induced to develop from isolated sorghum shoot apices (Bhaskaran et al., 1988). By manipulating plant growth regulators in the medium, sorghum shoot apices can give rise to not only single or multiple shoots but also embryogenic, shooty, or rooty callus (Bhaskaran et al., 1992). Other studies also revealed the versatile developmental patterns of sorghum shoot apices (Zhong et al., 1998), and that of shoot apices of maize (Zhong et al., 1992), rice (Christou and Ford, 1995), and oat (Avena sativa L.) (Zhang et al., 1996) under several different culture regimes. In turfgrass, shoot apex cultures were established for virus eradication and for long—term aseptic storage of germplasm (Dale, 1975, 1977a). Kinetin and 2,4-D (0.01-0.2 mg/L) in the culture medium were beneficial for plant regeneration from shoot apices (Dale, 1977b). The sizes of shoot apices excised were critical to virus elimination, survival rate in the culture, and regeneration behavior (Dale, 1975, 1977b). An interesting finding was that some 10 larger shoot apices produced an inflorescence instead of maintaining vegetative growth (Dale, 1975, 1977b). Besides vegetative shoot apices, floral apices of perennial and Italian ryegrasses were cultured for the production of functional pollen (Perez-Vicente et al., 1993). In this study, floral apices were excised from immature inflorescences and cultured in MS liquid medium supplemented with kinetin and 2,4-D to facilitate differentiation of spikelets. The differentiated spikelets completed full development on a hormone-free MS solidified medium. The shoot apical explant, derived from either vegetative or floral apex, could provide an alternative target for turfgrass transformation. In studies of turfgrass and cereal crops, plant regeneration from shoot apices was readily achieved and nearly genotype—independent (Sautter et al., 1995; Park et al., 1996). Plant regeneration from shoot apices can also be manipulated to avoid an intervening callus phase and adventitious regeneration, which increase the risk of undesirable somaclonal variation (Karp, 1994; Park et al., 1996). The competence of shoot apices was demonstrated for biolistics-mediated transformation of maize (Lowe et al., 1995; Zhong et al., 1996), rice (Christou and Ford, 1995), and for Agrobacterium-mediated transformation of rice (Park et al., 1996). Transient gene expression was also observed in meristem tissues of vegetative and floral apices of perennial ryegrass (Perez—Vicente et al., 1993) and wheat (Triticum aestivum L.) (Sautter et al., 1995). 11 Because shoot apex (or shoot meristems)—targeted transformation often has led to chimeric sectors in primary transformants (R0) (Christou and Ford, 1995; Sautter et al., 1995; Lowe et al., 1995; Park et al., 1996; Zhong et al., 1996), transgenic progeny can only be obtained from those transgenic sectors that happen to be germline cells in the shoot apices. In order to produce a high frequency of non- chimeric R1 transformants, follow-up manipulations on the sectored R0 transformants are often needed. The approach developed by Lowe et al. (1995) and Zhong et al. (1996) for maize should be seen as a solid step in this respect. By multiplication of shoot apices on the selection medium, non-I transgenic sectors were reduced in size or even eliminated as transgenic sectors enlarged and became predominant in each shoot apex. After these manipulations, transgenic sectors had a greater likelihood of contributing to the germline. Anther' Culture Haploids recovered from anthers have become an integrated part of modern breeding practices (Wenzel et al., 1995). The culture procedure is comprised of two steps: induction of microspore callus and subsequent regeneration. The induction medium contains an auxin, such as 2,4-D, indole-3—acetic acid (IAA), or naphthalene acetic acid (NAA), and cytokinin, such as kinetin; while the regeneration medium contains cytokinins, such as 6-BA. The later spikes with most of the microspores in the mid- or late-uninucleate stage, are 12 used to initiate the culture. Haploid plantlets have been regenerated from anther cultures of tall fescue (Kasperbauer et al., 1980); meadow fescue (Rose et al., 1987); Italian ryegrass (Boppenmeier et al., 1989); perennial ryegrass (Stanis and Butenko, 1984; Halberg et al., 1990; Opsahl— Ferstad et al., 1994a, b; Olesen et al., 1995b; Madsen et al., 1995), and the intergeneric hybrid (L. multiflorum x Festuca pratensis) (Rose et al., 1987). Genotype-dependence and inheritance of anther-culture response have been investigated (Opsahl—Ferstad et al., 1994a; Madsen et al., 1995). The genotype of perennial ryegrass determined the degree of correlation between different characters of androgenetic response, such as embryo-like structures per 100 anthers, and green plants per 100 anthers (Opsahl-Ferstad et al., 1994a). The high genotype-dependent anther response of ryegrass can be transferred via backcrossing to produce superior genotypes capable of anther culture (Halberg et al., 1990; Madsen et al., 1995). Strong genotype-dependence makes it necessary to modify the chemical composition of the culture medium and other in vitro conditions to fit the specific genotypes for improved androgenetic responses (Opsahl-Ferstad et al., 1994a). The total N content, ratio of N0{/ NHf, solid vs. liquid medium, and cold pretreatment affected the fate of anther culture in perennial ryegrasses (Opsahl-Ferstad et al., 1994b). 13 Effects of Genotype on In Vitro Regenerability The genotypic effect is a significant factor controlling the response of explants to in vitro culture conditions. For example, the competence for direct embryogenesis in orchardgrass was found to be genotype dependent (Hanning and Conger, 1982; Conger et al., 1983). The genotype regenerability dependency may be more pronounced in cell suspensions and protoplasts than in callus cultures (van Heeswijck et al., 1994), but the opposite situation was also reported (Olesen et al., 1995a). Genotype and culture response were correlated, including the correlation value of. callus induction—plant regeneration in tall fescue (Takamizo et al., 1990), meadow fescue (Wang et al., 1993a), and perennial ryegrass (Opsahl-Ferstad et al., 1994a; Olesen et al., 1995a). Genotypes with high regeneration percentages and high frequencies of green-plant recovery also showed superior suspension culture regeneration both in longevity and ability (Wang et al., 1995; Olesen et al., 1995a). Cultures derived from single genotypes may help to reduce inconsistent in vitro behaviors (Takamizo et al., 1990; Wang et al., 1995; Yamamoto and Engelke, 1997). This practice is more important for cool season turf species than warm—season species because most of the cool-season turfgrass species are genetically heterogeneous within cultivars or population (Huff, 1997). Single-genotype-derived cultures improved the efficiency of in vitro manipulations in heterogeneous populations (Takamizo et al., 1990; Wang et al., 1995), because researchers can 14 selectively use individual culture lines that have the best response to in vitro manipulations. Single-genotype-derived cultures from vegetative tissues of cross-pollinated or vegetatively propagated cultivars facilitate post-culture selection (Yamamoto and Engelke, 1997). This system could also help to evaluate and optimize other culture condition factors. Genotype selection within cultivars, especially cultivars with broad genetic backgrounds, for better response to in vitro cultures, would probably be effective for several turfgrass species, since androgenetic response in perennial ryegrass is probably determined by relatively few loci (Opsahl—Ferstad et al., 1994a). Transfer of culture response via backcrossing is a possible approach for non-culture responsive breeding material. Androgenesis superior to that of their parents was attained in hybrid clones of perennial ryegrass (Halberg et al., 1990; Madsen et al., 1995). 15 093— :8: co 215:8 $332.5 1:53“ , .H . _ . 1. .. .133, . ., .. :32 , ,. . .eunuscxv 5422 . . 13.959 «Econ—3.3:: .. ”1.1.0.93 «<5 .voo.....1_u._.m.._£§w3m$ 12.2.13 , 1131.11.11.5—831 _ 121.111.11.153 12. ., 1.1.11.5...» we :1 «o .553. EM. . £531.23— .18.:me b... unwa— 2% u .q1aoem56wmmm g3 .08.— .395093 9.3.3 _. _, .5 _ :8 3k... ©1133 .3335 five: awaken— amoa :1 .1... mi? .1182 .1 .o 933 smog ...1 10 93>» m2. 3.2315 333% name 58$ 1.1, 3 “:33. name .583 ..1 8 93>» 32 .g .1. 5 use? . g . 153533305. d. 2%— 2130024... E 82. 1.11.585 . 181111.315; .32 1.15.2 .25 .182. .1. .182. .w111531oa2 .1. 1. 11.11.3812 .52 2.5. 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A recent review by Lee (1996) listed six cool-season species of turfgrass transformed by a variety of transformation systems. The transgenes include the B-glucuronidase (GUS) gene, the phosphinothricin acetyltransferase (bar) gene, and the hygromycin phosphotransferase gene. Progress in turfgrass transformation has been made in exploring and optimizing transformation systems that have been used for other grass species and dicots. Agrobacterium-Mediated. Transformation Agrobacterium—mediated transformation has been widely used for many dicot plants as a convenient, highly efficient method for DNA transfer. A great benefit of Agrobacterium— mediated transformation is its capability to transform intact, regenerable plant tissues and organs (Horsch et al., 1985). Gene integration patterns are more predictable in Agrobacteriaum-mediated transformation than other gene transfer methods (Birch and Franks, 1991; Smith and Hood, 1995). Transformants from Agrobacterium—mediated transformation often contain genomic insertions by exact and single- or low- copy number of transgene cassettes (DeBlock, 18 1993). Single-copy insertions facilitate gene expression because they are less prone to cosuppression or gene- silencing effects that are usually found in multicopy insertion events (Vasil, 1995). Another benefit of Agrobacterium-mediated DNA delivery is the minimal exposure of explant material to tissue culture condition that induce genetic instability (Karp, 1994). Although grass and other monocots have been generally considered outside the host range of A. tumefaciens, Agrobacterium—mediated transformation has been used successfully in a few grass species including maize (Gould et al., 1991; Ishida et al., 1996), rice (Chan et al., 1992, 1993; Liu et al., 1992; Park et al., 1996), and wheat (Deng et al., 1988). An examination of successful monocot transformation points to several factors that may be critical to A. tumefaciens-mediated monocot transformation. These factors include the use of meristematic tissue for cocultivation with A. tumefaciens, the addition of phenolic compounds to the cocultivation media, and the use of the compatible strains of A. tumefaciens (see review by Smith and Hood, 1995). Agrobacterium—mediated transformation has not been reported in turfgrass. More efforts are warranted before this method becomes a routine transformation approach for grass species. 19 Transformation by Direct DNA Delivery DNA Uptake by Protoplasts Direct DNA delivery into protoplasts is one of the methods by which successful transformation has been achieved in turfgrasses. With the physical barriers of the cell walls removed, protoplasts can be induced to uptake DNA from the culture by osmotic treatment, e.g., polyethylene glycol, or electric shock (electroporation). Protoplast cultures have served as genotype-independent targets for free DNA uptake. By protoplast-mediated gene transfer, transgenic plants were” obtained from tall fescue (Wang et al., 1992; Ha et al., 1992; Dalton et al., 1995), red fescue (Spangenberg et al., 1994), orchardgrass (Horn et al., 1988), creeping bentgrass (Lee et al., 1995; Sugiura et al., 1997), and redtop (Asano et al., 1991; Asano and Ugaki, 1994). Although protoplast- mediated transformation has proven successful in these turfgrass species, plant regeneration from protoplasts is still difficult to achieve (Park et al., 1996) due to such uncontrollable parameters as genotype-dependent competence for regeneration (Potrykus, 1990; Hinchee et al., 1994; Spangenberg et al., 1995a). Other constraints of this process include infertility problems, undesired integration of multiple and rearranged transgene copies, and somaclonal variation resulting from the extensive cytogenetic disturbance in protoplast culture (Karp, 1994; Spangenberg et al., 1995a). 20 Biolistic Bombardment Microbiolistic delivery systems for turfgrass genetic engineering have achieved success. Invented by Sanford and others (Sanford et al., 1987; Sanford, 1988), biolistics delivers DNA molecules carried by tiny particles of Au or W into intact cells. Physical force generated from compressed H or gun powder to penetrate cell walls for DNA delivery circumvents the host range limitations of Agrobacterium vectors and also eliminates the need for plant regeneration from protoplasts. The biolistic DNA delivery method is versatile and culture-system independent, compared with protoplast DNA uptake methods. Several different types of explant material have been used successfully in turfgrasses to recover transgenic plants after biolistic bombardment. Transformation by biolistic delivery has been reported with embryogenic calli of creeping bentgrass (Zhong et al., 1993; Lee et al., 1995; Liu et al., 1998; Warkentin et al., 1997), and with cell suspensions of tall fescue and red fescue (Spangenberg et al., 1995a), creeping bentgrass (Hartman et al., 1994), and perennial ryegrass (Spangenberg et al., 1995b). Transgenic orchardgrass plantlets have been regenerated directly from leaf mesophyll cells after biolistic bombardment (McDaniel et al., 1997). Biolistic gene delivery also provides a unique way for direct transformation of organelle genomes and rapid assessment of transient expression of genetic constructs in plants (Birch and Franks, 1991). 21 Although biolistic DNA delivery methods were suggested to be universal for every species (Birch and Franks, 1991), they have the shortcomings analogous to those in other direct gene transfer methods. The complex gene integration patterns that often result from this method may have significant implications for developing stable transgenic plants. Microinjection The production of transgenic plants from microinjected cells has only been successful in tobacco (Nicotiana tabacum L.) and rapeseed (Brassica napus L.) (Neuhaus et al., 1987). The microinjection apparatus consists of three basic components: a micropipette made of glass, an injection system applying injection driving force to the micropipette, and a micromanipulator controlling the microscopic motions of the injection pipette (Mathias, 1987). Since it can deliver a DNA solution into the nuclei of the target cells, microinjection has the potential to transform any plant species when the proper target cells are available (Hinchee et al., 1994). The potential target cells for whole-plant transformation by microinjection were listed by Mathias (1987). Among these cells, protoplasts, suspension culture cells, and callus cells are the most readily available turfgrass in vitro cultures and therefore have the potential to become the target cells in microinjection-mediated transformation. Nevertheless, the feasibility of using microinjection for whole-plant transformation is restricted by the technical 22 difficulties. The operation of microinjection is laborious and requires the difficult process of regenerating plantlets from isolated single cells or small cell clusters because large tissues cannot be resolved clearly enough for accurate needle placement (Hinchee et al., 1994). This task is made even more demanding when protoplasts are used as the target cells in transformation, since the plating efficiency and regeneration efficiency from protoplasts also play a significant role in determining transformation efficiency (Hinchee et al., 1994). Compared with other gene delivery systems, microinjection is not as well proven for creating transgenic plants. Turfgrass transformation via microinjection has not been reported. Silicon Carbide Fiber—Mediated Transformation Silicon carbide fiber-mediated transformation for plant cells was developed by Kaeppler et al. (1990). This method involves vortexing the cell or tissue culture with plasmid DNA and CSi fibers averaging 0.6 pm in diameter and 10—80 pm in length, which facilitate DNA delivery into nuclei by serving as microinjection needles (Kaeppler et al., 1990, 1992). Progress in CSi-mediated DNA delivery has been slow. The scarce number of studies include earlier reports on cell suspensions of maize, tobacco (Kaeppler et al., 1990), redtop grass (Asano et al., 1991), and a more recent one using mature embryos of wheat (Serik et al., 1996), in which only GUS transient expression was detected in the CSi-DNA-treated 23 cultured cells. Stable transformation of plant cells was first reported by Kaeppler et al. (1992) in maize. The study by Frame et al. (1994) was the first and the only one so far, in which fertile, transgenic maize plants were recovered from plant cells transformed via CSi-mediated gene delivery. The transformation frequency was estimated as 10'6 transformants per cell treated (Kaeppler et al., 1992), lower than with biolistic bombardment. Nevertheless, it might be premature to judge this method solely on the transformation frequency data generated from one or two studies, since the transformation frequency of maize cells using other methods varied significantly from case to case with differences up to two magnitudes, as shown by Kaeppler et al. (1992). The observed reduction in cell viability as the result of CSi treatment (Kaeppler et al., 1992) is a potential drawback of this method. Advantages of Csi fiber—mediated transformation over biolistic bombardment include very low equipment cost, relative ease of the procedure (Kaeppler et al., 1992), and the better control of DNA quantity delivered (Thompson et al., 1995). While these features are attractive to plant researchers, the potential of this method needs to be further explored in order to obtain transgenic plants in other species. Regenerable targets of gene delivery should be extended to include tissues and organs other than suspension cells. 24 Selection for Transgenic Plants Cells containing gene(s) of interest are selected and regenerated into individual plants. A selectable marker gene in transgene constructs is usually necessary to confer the preferential growth of transformed cells in the presence of the corresponding selective agent. Except for the case in which creeping bentgrass was transformed by Zhong et a1. (1993) using the GUS gene as the screening marker, all other transformation events in turfgrass species have relied on using selectable markers for identifying transgenic plants. Antibiotic markers, such as hygromycin resistance, and herbicide markers, such as phosphinothricin resistance, have been successfully transferred and expressed in transgenic turfgrasses as selectable markers (see the list of selectable markers in the review by Lee, 1996). Selection schemes used for turfgrasses vary widely among different genotypes and culture types in terms of concentrations of selective agents and the timing of application because of the differing sensitivity of cultures to the selective agents. Since selection efficiency can be measured as a function of the recovery rate of transgenic cells and the non—transgenic “escapes”, the minimal lethal dosage of the selective agents to non-transgenic cells must be empirically determined. Selection by herbicides usually needs to be delayed for a certain period of time after gene transfer in order to allow the transformed cells to recover from physical damage (especially in electroporation and particle bombardment) and 25 to produce sufficient quantities of the resistance enzyme for protection. Either a fixed concentration or stepwise increase in concentration of selective agents has been used frequently in the selection schemes for turfgrass transformation. The finding that continuous vs. discontinuous selection schemes are correlated with the copy number of integrated transgenes in tall fescue (Dalton et al., 1995) might be important in efforts to minimize down-regulating or “gene-silencing" effects caused by multiple integrations. Selectable markers are essential to genetic transformation. However, fewer selectable markers can be used for monocot species than for dicot plants (Zhou et al., 1995). Given the fact that the presence of a selectable marker gene in plant genomes makes it impossible to carry out the subsequent gene transfers with the same marker, it would be beneficial to develop strategies to remove selectable markers from the transgenic plants. Transformation by cotransferring unlinked transgenes has been achieved with high efficiencies in several crops, including maize (Schocher et al., 1986; Gordon-Kamm et al., 1990), rice (Chair et al., 1996), barley (Wan and Lemaux, 1994), rapeseed (Herve et al., 1993), and tobacco (Carrer and Maliga, 1995). This method was originally used to speed up transformation because it eliminates the need to prepare a transgene construct harboring both the desired gene and the selectable marker (Schocher et al., 1986). It also provides a simple approach to recycling selectable markers. If there is 26 no physical linkage between the selectable marker and the desired transgene, removal of the selectable-marker locus will be readily achieved by sexual segregation. However, the cotransferred genes could become linked in the genomes of the transformants if they are co-precipitated and transferred by using the biolistic method (Gordon-Kamm et al., 1990; Spencer et al., 1990). Another strategy for removing selectable marker genes involves using site-specific recombination systems obtained from microorganisms, such as budding yeast (Saccharomyces cerevisia) (Cregg and Madden, 1989), fission yeast (Schizosaccharomyces pombe) (Lyznik et al., 1993), and bacteriophage P1(Dale and Ow, 1991; Sauer, 1994). Each of these systems consists of a pair of inverted repeat DNA sequences as the recognition site and a specific recombinase gene. The specific enzymatic activity of this recombinase results in the excision of the inverted repeats and the sequence between them. Plasmid constructs for gene transfer can be made to contain the selectable marker flanked by the inverted repeats and used to transform plant cells. As the established recombination target, the selectable marker can be excised from the plant genome by the corresponding site— specific recombinase activity, which can be introduced through either transfer of the recombinase gene or direct application of purified recombinase enzyme (Dale and Ow, 1991). Biolistic bombardment systems may be suitable for these manipulations. 27 Recycling of selectable marker genes should be included in transformation practices in turfgrass species. Although the strategy of using recombination systems has not yet been tested in turfgrass, there is strong reason to be optimistic about its success, since gene removal has already been achieved in tobacco by the Cre/lox recombination system (Dale and Ow, 1991), and the yeast FLP/FRT recombination system has already been tested to be functional in maize and rice protoplasts (Lyznik et al., 1993). After the fertile transgenic plants have been regenerated, selfing is usually a necessary step to obtain homozygous seed progeny from the heterozygous parents at the transgene loci. Self-fertilized turfgrass species provide a more convenient system to achieve this task than the outcrossing species, in which self—infertility limits inbreeding, and apomictic species, in which either self- fertilization or cross-fertilization is limited. Surrogate Transformatin via Grass Endophytes Endophytes are organisms that are contained or grow entirely within plants and spend all or nearly all of their life cycles in their hosts (Siegel, 1987; Breen, 1993a, b). Among grass endophytes, true endophytes (endophytes sensu stricto, Schardl, 1994), are currently recognized as most relevant in grass breeding. This type of endophyte never produces external reproductive stages on the host plants and thus is less destructive and even relatively symptomless to the hosts compared with those that produce external mycelium or spores. In fact, symbiotic associations between true endophytes and grass hosts could be characterized as stable and mutualistic; the endophytes are transmitted asexually via host seeds. Manipulating the performance of grass hosts via genetic engineering of the associated endophytes represents an alternative approach to transform grasses (Murray et al., 1992). Surrogate transformation has been successful by transforming Acremonium spp. (Murray et al., 1992; Tsai et al., 1992), and the reintroduction of transformed Acremonium into perennial ryegrass (Murray et al., 1992). Progress in transforming filamentous fungi (reviewed by Fincham, 1987) using protoplasts along with polyethylene glycol [a-hydro-m- hydroxypoly(oxy-1,2-ethanediyl)] treatment and osmotic conditioning provides the basic protocol for grass endophyte transformation. Transformation frequencies for Acremonium spp. were achieved up to 700 to 800 transformants ug'1 plasmid DNA (Murray et al., 1992), depending on such parameters as regeneration medium (Oliver et al., 1987; Murray et al., 1992), osmoticum techniques, and states of vector DNA (linear or circular) (Murray et al., 1992). Electroporation was used by Tsai et al. (1992) to facilitate DNA transfer because it requires few post-manipulations and might facilitate homologous integration or gene conversion (Schardl, 1994). Surrogate transformation makes it possible to manipulate grass hosts by introducing heterologous genes into endophyte 29 genomes. Most known molecules responsible for endophyte- enhanced plant protections are the end products of biosynthetic pathways (Schardl, 1994). Examples are peramine, which confers protection to the hosts from damage by Argentine stem weevil (Rowan et al., 1986), and lolitrems, the cause of ryegrass staggers syndrome (Gallagher et al., 1981, 1982, 1984; Weedon and Mantle, 1987). Identification of the genes responsible for rate-limiting steps would be very critical in attempts to manipulate endophytes. Progress has been made with other filamentous fungi (Schardl, 1994). Dimethylallyltryptophan synthase (DMAT synthase), which is involved in ergoline alkaloid biosynthesis pathways, was isolated from Claviceps purpurea Fr. (Tul.) (Shibuya et al., 1990). Because of possible shared homologies due to the evolutionary relatedness in endophytes, genetic studies of turfgrass endophytes would certainly benefit from findings in other filamentous fungi. By using state-of-the art molecular techniques as listed by Schardl (1994), various gene manipulations might be readily performed for grass endophytes once the relevant genes have been cloned and characterized. In addition to improving turf and forage grasses, endophyte strategies could be employed to implement molecular farming to produce an array of chemicals such as natural plant regulators, insecticides, natural herbicides, and medicinal agents (Porter, 1994). 30 FUTURE OPPORTUNITIES IN TURFGRASS BIOTECHNOLOGY Turfgrass has a great impact on people’s lives in the United States. The cash value of the turfgrass industry had grown from 4 billion dollars in 1974 to 45 billion dollars in 1994 and will approach 90 billion dollars by the end of this century (Duble, 1996). However, the turfgrass industry faces great challenges, including water shortage, rising costs of energy, limited labor resources, and environment—oriented restrictions on the use of fertilizers, herbicides, fungicides, and pesticides (Duble, 1996). Biotechnology could play an important role in providing technical solutions to these problems. Previous progress has proven that the necessary elements of applying biotechnology to turfgrasses have become available in practice, and it is feasible to use biotechnology to solve certain problems in turfgrasses (van Heeswijck et al., 1994). With genetic engineering as the central element (Boulter, 1995), biotechnology provides a set of powerful tools in the development of desirable turfgrass cultivars, including cultivars tolerant to drought and salts for water conservation, cultivars with resistance to microbial diseases and insect pests, and herbicide-resistant plants that can be used to simultaneously control both weeds and turfgrass diseases after being sprayed with glufosinate herbicides, which were found to have an inhibition effect on fungal diseases (Liu et al., 1998). Other desirable tasks are to develop turfgrass cultivars with low maintenance 31 requirements such as dwarf or rosette turfgrass (Xu et al., 1995). Nevertheless, the potential of biotechnology in turfgrass improvement largely depends on how well plant regeneration and gene expression is understood as well as availability of the new genes of important functions. The recent progress in functional genomics could take biotechnology of turfgrass a big step forward. Based on technologies of miniturization (DNA chips or microarrays) and use of computers (data analyzing, storing, and retrieving), the high-throughput methods accelerate geneomic sequencing and the discovery of gene functions. As the direct result, gene cloning will also be greatly accelerated by the information from the complete genomic sequences of the model plants such as Arabidopsis, which is expected to be finished by the year 2001 (Bouchez and Hofte, 1998). Turfgrass biotechology will see the growing number of genes that can be used for turfgrass genetic engineering. Traditionally, funding by government and large industry for turfgrass research has been and will continue to be limited. The progress in turfgrass biotechnology will greatly benefit from achievements in other grass species. Examples of achievements in other grass species include the development of somaclonal variants that tolerate environmental extremes, such as salt—tolerant rice (Bong et al., 1996), chilling- tolerant rice (Bertin et al., 1996), drought-tolerant rice (Adkins et al., 1995), frost-tolerant wheat (Sutka, 1994), drought-tolerant wheat (Hsissou and Bouharmont, 1994), dwarf 32 wheat (Guenzi et al., 1992), glyphosate-tolerant barley (Escorial et al., 1996), drought- and acid- soil— tolerant sorghum (Waskom et al., 1990), eyespot disease [Pseudocercosporella herpotrichoides (Fon) Deighton]- resistant sugarcane (Saccharum officinarum L.) (Ramos et al., 1996), Fusarium -resistant wheat (Ahmed et al., 1991), leaf rust (Puccinia recondita Roberge ex Desmaz.)-resistant wheat (Oberthur et al., 1993), and downy mildew [Sclerophthora macrospora (Sacc.) Thirum, Shaw, and Naras.]—resistant millet [Pennisetum glaucum (L.) R. Br.] (Nagarathna et al., 1993). 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Brown, C.G. Santhino, and J.E. Fry. 1995. Glyphosate-tolerant CP4 and GOX genes as a selectable marker in wheat transformation. Plant Cell Rep. 15: 159-163. 58 CEBU?TERLEHWO CHITINASE GENES AND PLANT DISEASE RESISTANCE (Literature review) INTRODUCTION Agricultural crops suffer from a vast array of diseases caused by microbial attack. Fungal diseases are among the major factors limiting crop productivity and incur huge capital expense on using conventional pesticides, which are at possible environmental cost. Nevertheless, plants have 1 evolved mechanisms to protect themselves from invasion by fungal and other pathogens. Plants react to pathogenic invasion by the coordinated activation and expression of a number of genes. Many of these genes encode enzymes involved in synthesis of products such as phytoalexins and lignin that impede or inhibit the infection. Other genes encode proteins that are assumed to act directly on the invading pathogens. Genes that encode certain pathogenesis-related (PR) proteins such as chitinase fall in this category. 59 CHITINASES IN PLANTS Chitinases are proteins capable of hydrolyzing the fi—l,4 linkage of the N-acetyl-D-glucosamine (NAG) polymer chitin [poly [1,4—(N-acetyl-B—D-glucosaminide)]]. A plant chitinase was first purified from wheat in 1979 (Molano et al., 1979). Since then, Chitinases have been found in more than 30 plant species (Graham and Sticklen, 1994). Up to now, the plant species from which chitinase genes that have been identified and sequenced include Arabidopsis thaliana, tobacco (Nicotiana tabacum), bean (Phaseolus vulgaris), wheat (Triticum aestivum), potato (Solanum tuberosum), rice (Oryza sativa), maize (Zea mays), barley (Hordeum vulgare), oilrape (Brassica napus), grapes (Vitis Vinifera), cotton (Gossypium hirsutum), sugar beets (Beta vulgaris), cacao (Theobroma cacao), poplar (Populus trichocarpa), common ice plant (Mesembryanthemum crystallium), Arabis lyratasuch (National Center for Biotechnology Information, website: www.nobi.nlm.nih.gov). Most plant Chitinases isolated are endochitinases, i.e. they degrade chitin from hydrolysis sites within the polymer, and produce fragments two to five NAG residues long (NAGkS) (Koga et al., 1989) as the result of the preference for the second or third fi-1,4 linkage from the polymer’s reducing end. Many plant Chitinases appear to be organ-specific and developmentally regulated. For instance, the tomato Class II 60 endochitinase encoded by the Chi2 gene is predominantly expressed in pistils and is temporally regulated (Harikrishna et al., 1996). Nevertheless, physiological functions of plant Chitinases are still unknown (Graham and Sticklen, 1994). Current interest in Chitinases arose from the correlative evidence that they are part of plant defense mechanisms. Many Chitinases are among the so-called pathogenesis-related (PR) proteins (Linthorst, 1991; Graham and Sticklen, 1994). Concentrations of the PR proteins are usually maintained at low constitutive levels in optimal growth conditions but can escalate to much higher levels upon wounding, pathogenic challenges such as infection of the plant tissue with fungal or bacterial pathogens, or by biotic and abiotic challenges and elicitation such as from fungal or plant cell wall material, or from ethylene (Majeau et al., 1990; Roby et al., 1990). Many Chitinases have been shown to have anti-fungal activity. They are capable of catalyzing degeneration of chitin—containing fungal cell walls and consequently inhibit fungal growth at hyphal tips (Boller et al. 1983; Mauch et al. 1988; Schlumbaum et al., 1986; Arlorio et al., 1992; Gomes et al., 1996). Nevertheless, the roles of chitinases in plant defense systems are not limited to their lytic activity. The findings by Takemoto et a1. (1997) with a basic chitinase in potato suggest that Chitinases may be involved in cytoplasmic aggregation, an early resistance-associated event, because of the chitinase ability to bind actin. 61 Numerous studies have been focused on characterization of the chitinase enzymes from various plant species, their regulation, localization, and physiological roles in plant defense systems against fungal pathogens. A recent trend is the shifting towards the development of transgenic crop plants with increased levels of expression of chitinase genes in hopes of generating fungal disease resistant varieties. ENGINEERING FUNGAL RESISTANCE USING CHITINASE GENES Traditional breeding has been employed to strengthen the endogenous defense capacity of plants by introducing antimicrobial genes into the germplasm through crossing and selection. Gene introgression using traditional methods is, however, confined to individual gene pools, in which natural gene flows occur. Breeding for microbial resistance by traditional crossing and selection is also restrained by genetic linkages, which impede or even prevent single gene introgression. An emerging technology to lift the restraints on traditional breeding methods is based primarily on introducing and expressing single antimicrobial genes in a constitutive manner. As suitable transformation systems become available and as more antimicrobial genes have been isolated, cloned, and characterized, this approach has become a practical option. Genes encoding Chitinases are among the 62 most attractive genes for this approach because of their lytic activity against fungal pathogens (Lamb et al., 1992). Transgenic plants with a chitinase gene were first developed in tobacco by Broglie et al.(1991). Bioassays showed that the homozygous transgenic tobacco plants expressing high levels of chitinase grew faster, had a lower seedling mortality rate, and suffered less root weight loss than the untransformed plants when they were challenged with the soil-borne pathogen Rhizoctonia solani. Chitin lytic activity was one of the roles that Chitinases played in conferring resistance on transgenic plants. The bean chitinase expressed in canola plants appeared to account for breakdown of cell walls of the invading R. solani in vivo (Benhamou et al., 1993). This study marked the beginning of research for engineering pathogen resistance using chitinase transgenes. Since then, a number of studies have been published on the efficacy of Chitinases in enhancing disease resistance, mixed results have been obtained from these studies in terms of the relative effectiveness of Chitinases in increasing disease resistance. Improved disease resistance conferred by chitinase genes was reported in transgenic plants of a number of species, including oilrape (Brassica napus var. oleifera, Grison et al., 1996) expressing a tobacco chitinase gene, cucumber expressing a rice chitinase gene (Tabei et al., 1998), rose (Rosa hybrida L.) expressing a rice chitinase gene (Marchant 63 et al., 1998). Samac and Shah (1994), on the other hand, supported the importance of chitinase in plant defense when they observed the increased susceptibility to fungal pathogen Botrytis cinerea in transgenic Arabidopsis thaliana plants transcribing antisense RNAs to the Class I chitinase gene. The differential antifungal activities of different isoforms of Chitinases have been recognized. In the study by Punja and Raharjo (1996), transgenic carrot expressing the tobacco chitinase gene exhibited significant improvement in disease resistance against Botrytis cinerea and Rhizoctonia solani, while transgenic cucumber expressing the same genes did not show improved resistance. This study also showed that expression of petunia chitinase genes in tobacco or carrot plants did not enhance their resistance to Botrytis cinerea and Rhizoctonia solan. Studies by Sela-Buurlage et a1, (1993) aslo showed the differential antifungal activities of other tobacco chitinase. These studies, in general, demonstrated the efficacy of expressing chitinase gene to confer disease resistance. Overall, the general effectiveness of chitinase transgenes was inconsistent when plant species, type of chitinase protein expressed, and the characteristics of the fungal pathogen tested were considered. In search of more effective and lasting resistance, co- expressions of multiple antifungal proteins were attempted. When compared with “isogenic” transformants expressing a single gene, Jach et al. (1995) found that higher levels of protection against Rhizoctonia solani were acquired in tobacco plants transformed by the tandemly linked genes encoding a Class II chitinase, a Class II B-1,3-glucanase, and a Type-I ribosome-inactivating protein from barley. They suggested that this “multigene” tolerance be due to synergistic interactions among the co-expressed antifungal proteins. Lamb’s group (Zhu et al., 1994) convincingly demonstrated synergistic interactions between chitinase and B-1,3-glucanase. In their study on tobacco transgenic plants, disease resistance conferred by the combination of chitinase and glucanase was significantly more effective than either transgenes alone at the double dosages. Transgenic approaches have been used to study chitinase expression patterns in response to inducing factors. In these studies, transgenic plants were made with chimeric genes, which were the fusions of promoters of various chitinase genes to the coding sequence of a reporter gene. Xu et al. (1996) introduced the rice chitinase promoterzzGUS (B- glucuronidase) fusion into rice plants and demonstrated that the 5'-flanking region of the rice basic chitinase gene RC24 was stress-responsive and tissue specific in activating the transcription of RC24. Kellmann et al. (1996) confirmed in transgenic tobacco that genes encoding two class II Chitinases from peanut (Arachis hypogaea L. cv. NC4) responded differently to ethylene, salicylate, or fungal spores. An ethylene-responsive region was identified in tobacco Class I chitinase CHN48 to contain two copies of 65 GCC-box as a regulatory enhancer for gene transcription (Shinshi et al., 1995). The fact that this GCC-box has been found conserved in a number of ethylene-responsive defense genes suggests possible coordinations among defense mechanisms. In other studies, Fukuda et a1. (1994) and Fukuda (1997) identified two sequence repeat sites in promoter element ElRE of the tobacco Class I chitinase gene CHNSO to be the motifs involved in binding of the nuclear protein factors. Wounding activation of the poplar chitinase was also confirmed by using chimeric constructs of chitinase gene promoters and the reporter gene (Clarke et al., 1994). Promoter studies of Chitinases revealed important relationships between binding of elicitors to the promoter regions and the induction of gene transcription. Silencing of chitinase genes was also documented in transgenic plants. Two related studies (Kunz et al., 1996; Schob et al., 1997) reported that a high incidence of gene silencing by homologous copies of tobacco Class I chitinase A gene in transgenic tobacco plants. They concluded that silencing depended on transcribed sequence homology and the transgene dosage and that 60% identity of sequences encoding the catalytic domain was required for silencing. Their results further suggested that chitinase gene silencing in these cases was post-transcriptional, because the levels of steady—state mRNA in the “silenced" plants decreased greatly, while the levels of nascent mRNA were the same as in the highly expressing plants. The study by Hart et al. (1992) 66 also suggested that co-suppression instead of cytosine methylation of the transgenes resulted in silencing of a homologous chitinase gene (Class I) in transgenic tobacco. Other important conclusions from this study were that gene silencing was organ position-dependent, potentially reversible, and of non-Mendelian inheritance. Chitinase genes from non-plant organisms such as insects, fungi, and bacteria have also been used in developing pathogen resistance in plants. In a study by Terakawa et al. (1997), Chil, a chitinase gene involved in cell autolysis of the filamentous fungus Rhizopus oligosporus, was found to suppress infections by the discomycete pathogens Sclerotinia sclerotiorum and Botrytis cinerea in transgenic tobacco plants. Transgenic plants that constitutively expressed hornworm chitinase were produced and found to have higher resistance to insect pests (Kramer and Muthukrishnan, 1997). Evidence supports that chitinases are an integrated part of the natural mechanism whereby the plant responds to pathogen infections. The strong correlation between chitinase enzymatic activity and its role in conferring resistance to filamentous fungal pathogens makes chitinases the good transgenes for developing host resistance in plants using genetic transformation approach. 67 C3UUNEER.ITHUME GENETIC TRANSFORMATION OF CREEPING BENTGRASS (AGROSTIS RALUSTRIS HUDS.) USING A ELM CHITINASE GENE ABSTRACT An effective microprojectile-mediated transformation system was used to produce transgenic creeping bentgrass' (Agrostis palustris Huds.) plants with the elm chitinase(H52) gene (Sherald et al., 1994). The selectable marker bar (phosphinothricin acetyl-transferase, PAT) gene and the nonselectable H82 gene on separate plasmids were co-introduced into the creeping bentgrass genome, when embryogenic calli of creeping bentgrass were bombarded with tungsten particles coated with DNA of both plasmids. Transgenic plants carrying the H82 gene were identified from bar gene-transformed plants, which were resistant to the herbicides bialaphos and Ignite““..A total of eleven transformed lines carrying the H82 gene were obtained from forty-seven independent transgenic lines carrying bar gene. The frequency of the unlinked cotransformation of bar gene and the H82 gene is 23.9%, which was comparable to that of 68 other studies on different plant species and transgenes using unlinked co-transformation approach. The expression of the H82 gene was detected from some transgenic lines of creeping bentgrass at both the transcriptional and the translational levels. 69 INTRODUCTION Like other field-grown crops, turfgrass in golf courses is continually endangered by fungal pathogens. Close mowing, frequent irrigation and constant bruising from traffic make golf greens even quite vulnerable to attack by fungal pathogens. Hence, synthetic fungicides are commonly and frequently applied to golf courses to provide continued control of damage caused by pathogenic fungal populations. It was estimated that the turfgrass industry alone spent over S 80 million yearly for chemical fungicides (Vargas, 1994). The extensive and heavy reliance on fungicides not only adds to the operational cost of turf industry, but also arouses the legitimate concerns over the hazard those fungicides pose to the environment. The stakes are so high that alternatives strategies have been sought for protecting golf turf (USGA, 1997). Creeping bentgrass is a major cool season species used for golf greens, tees, and fairways. This important turf species is susceptible to a variety of fungal diseases such as brown patch (Rhizoctonia spp.), dollar spot (Sclerotinia homoeocarpa), and Pythium blight (Pythium spp.) (Smiley, 1983). Using transformation technology to strengthen endogenous resistance to these pathogens is not only a desirable but also an achievable objective, for 70 transformation systems have been developed and are being continually improved. In this chapter, I describe the transformation and regeneration of stably transformed creeping bentgrass plants by using a Biolistic”‘nediated transformation system on embryogenic calli and a cotransformation strategy to efficiently recover transgenic plants with the elm chitinase gene (H32) insertions. Transgenic plants were analyzed for gene integration and expression. Some transgenic lines were found to have enhanced (up to 3-fold increase) resistance to the turfgrass disease brown patch (Rhizoctonia solani) by our research collaborators. 71 MATERIAL AND METHODS Plant Material Creeping bentgrass (Agrostis palustris Huds.) is an allotetraploid (2n=4x=28), cool season species widely used in golf courses. The cultivar “Penncross” (Penn State University), which is a synthetic cultivar produced by the random crossing of three vegetatively propagated clones (Hein, 1958), and “Putter” (The Scotts Company, Marysville, Ohio) were used as the target plants for microprojectile- mediated transformation. The Transgenes and‘Vectors Two plasmids, pKYLX71-HSZ carrying the elm chitinase gene (H82) and pJSlOl carrying bar gene and the mannitol-l- phosphate dehydrogenase gene (mtlD) were used for transformation. pJSlOl was constructed and kindly provided by Dr. Ray Wu of Cornell University. pKYLX71-H82 was constructed by Mr. Don Warkentin of Turfgrass and Cereal Genetic Engineering Laboratory at Michigan State University. Both bar and the H82 genes were driven by the cauliflower mosaic virus (CaMV) 358 promoter, while mtlD gene was driven by the rice actin 5’ promoter (McElroy et 72 al., 1991). Diagrams of each plasmids with major restriction sites are shown in Figure l. Inducing Embryogenic Calli from.Mature Seeds Embryogenic calli were initiated from mature seeds (caryopses) following the protocol developed by Zhong et al. (1991). Seeds were first surface-sterilized in aqueous solution containing 50% (v/v) of commercial grade CloroxG) (active ingredient 5.25% NaClO), 0.1% Tween 20 (polyoxyethylene-sorbitan monolaurate), and 0.02% HgCl, and stirring under vacuum for 15 minutes. The seeds were then soaked in 70% ethanol for 5 minutes and rinsed in a large volume of sterilized water three times before being transferred to the autoclaved semi—solidified callus induction medium containing MS (Murashige and Skoog, 1962) basal salts and vitamins (Sigma Chem. Co. 5519), 0.5 g/l enzymatic casein hydrolysate, 3% (w/v) sucrose, 30 DM dicamba (3,6-dicloro-o-anisic acid), 9 HM BA (6- benzyladenine), and 7 g/l of phytagar (Gibco BRL Co.). Cultures were incubated at 25°C in the dark for one month before embryogenic calli appeared from coleoptiles. Only the friable and yellowish calli were selected for the subculture and multiplication. Subculture was carried out using the same calllus induction medium. Callus derived 73 from individual seeds was kept separately to ensure initial genetic homogeneity within each callus line. For calli on the solid medium, a 2-week interval was used for subculture. Suspension cultures were initiated by transferring 2 g of friable calli (one month old) into a 225 ml flask containing 50 ml of the liquid callus induction medium and incubated at 25°C in dark with shaking at 150 rpm. Half of the medium was removed and replenished with the same volume of fresh callus induction medium every three days. Each flask of suspension culture was divided| into two cultures every nine days. microprojectile-mediated Gene Delivery Friable and yellowish embryogenic calli were selectively collected and placed to form a thin layer of about 2 cm in diameter on the central surface of the solidified callus induction medium in a 15 x 60 mm Petri dish. In the cases cell suspensions were used, a piece of #2 Whatman filter paper (5.5 cm in diameter) was placed on the central surface of the medium and the 2 ml of the suspension culture (about 0.2 g of cells) were pipetted onto the center of the filter paper, and then the excess liquid removed. M 17 tungsten particles (0.9-1.20m in diameter, Sylvania, Towanda, PA) were used as 74 microcarriers, which were prepared in aliquots with 100% ethanol at a density of 30 mg tungsten per ml ethanol. The preparations were treated with a probe sonicator to break the aggregates of tungsten particles. Plasmid DNA was precipitated onto tungsten particles with CaClzand spermidine following a modified protocol from Klein et al.(1987) after the tungsten particles were washed twice with sterile distilled water in a microcentrifuge tube. One hundred ul of the final microcarrier suspension (containing three mg of the tungsten particles) was transferred into a microcentrifuge tube and vortexed continuously while pipetting aliquots of the suspension to avoid non-uniform sampling. A total amount of 25 pg of the mixed plasmid DNA (pJSlOl and pKYLX71-H82) at the ratio of 1:1 (molar:molar), 100 ul of 2.5 M CaCl2 and 40 ul of 0.1 M spermidine (free base, tissue culture grade, Sigma Chemical Co.) were sequentially added into the microcarrier suspensions during continuous vortexing. The microcarrier-DNA coating mixture was vortexed for another ten minutes. DNA—coated microcarriers were collected at the bottom of the microcentrifuge tube by centrifuge after the supernatant was decanted, and then washed with 250 ul of 100% ethanol twice using vortexing and centrifuging. DNA-coated microcarriers were re-suspended in 100 ul of 100% ethanol. 75 Biolistic® PDS-lOO/He system (Bio-Rad Laboratories, Inc.) was used for gene delivery. Ten ul of DNA-coated tungsten suspension was pipetted onto the center of each macrocarrier, which was serving as the vehicle for the tungsten particles. After the ethanol was dry, the macrocarrier was loaded on the system for bombardments, which were then carried out at callus or suspension preparations. Each callus or suspension preparation was bombarded three times within 5 minutes. GUS transient expressions per plate were used as the reference to adjust parameters of the system for maximizing gene delivery efficiency. The standardized bombardment protocol was as followed: rupture disc-pressure: 1,550 psi; gap-distance from rupture disc to macrocarrier: 6 mm; macrocarrier travel distance: 16 mm; and microcarrier travel distance: 6 cm. Selection Regime Bialaphos (L-phosphinothricinyl—L-alanyl—L-alanine) (provided by Meiji Seika Kaisha of Japan), a tripeptide consisting of one L—phosphinothricin (PPT) and two D- alanine residues, was used as the selective agent for selecting the transformed cultures expressing the bar gene. Uptake of bialaphos by plants or bacteria results in 76 cleavages between the residues by intracellular peptidases and the subsequent release of L—PPT, which is a potent inhibitor of glutamine synthetase, an important enzyme in ammonium assimilation pathways. Loss of normal activity of glutamine synthetase leads to the accumulation of ammonium, which eventually kills cells because of its toxicity. Transgenic cultures expressing bar gene would be able to synthesize phosphinothricin acetyltransferase (PAT), which detoxify L—PPT, in the presence of acetyl-CoA, by catalyzing acetylation of PPT into a nontoxic product, N- acetyl—PPT. To ensure the effective recovery of transgenic plants from the majority non-transgenic plant population, callus induction medium containing bialaphos at the concentration, 5 mg/L and 10 mg/L, and the regeneration medium containing bialaphos at 15 mg/L, were used over the sixteen weeks of in vitro selection. Callus initiated from each seed was maintained separately to ensure initial genetic homogeneity within callus lines. Negative control was made by bombarding embryogenic calli with tungsten particles without any plasmid DNA. Three days after the bombardment, initial selection began with calli subcultured onto the callus induction medium that contains 5 mg/L of bialaphos. Callus and the subsequently regenerated, rooted plantlets from the same 77 surviving piece of callus in the first selection cultures are defined as a line. Regeneration of plantlets was carried out for 4 weeks on the solidified regeneration medium containing MS (Murashige and Skoog, 1962) basal salts and vitamins (Sigma Chem. Co. 5519), 3% (w/v) sucrose, 0.7% phytagar, and 15 mg/L of bialaphos. DNA Isolation and Analyses A modified C-TAB method (Rogers and Bendich, 1985) was used to isolate high molecular weight DNA from nontransgenic as well as putative transgenic plants. Twenty grams of fresh leaf tissue was chilled in liquid nitrogen and ground into fine powder in mortar and pestle. The leaf material was transferred into a 250 ml oakridge tube and incubated at 60°C in a water bath for two hours with 70 ml of the C-TAB extraction buffer (0.1 M Tris—HCl, 1.4 M NaCl, 0.02 M EDTA, 2% C-TAB (hexadecyltrimethylammonium bromide, 1% polyvinyl-pyrrolidone (mol. wt. 40,000, Sigma Chemical Co.), and 1% 2-mercaptoethanol). To the cooled extraction mixture, 70 ml of chloroform/isoamyl alcohol (24:1) was added, and incubated for 20 minutes at room temperature. The upper, clear aqueous phase containing DNA was separated from the cell debris in the bottom phase and transferred to 78 a clean oakridge tube after centrifugation for 10 minutes at 10,000 g. The aqueous phase was then extracted twice with chloroform/isoamyl alcohol (24:1) and twice with phenol/ chloroform/isoamyl alcohol (24:24:1). DNA was spun down into a pellet for 10 minutes at 5,000 g after 1/10 volume of 3 M sodium acetate and 0.75 volume of isopropanol were added to the aqueous portion, and chilled for two hours at -20°C. DNA pellets were dissolved in 0.5 ml TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) and transferred to a microcentrifuge tube. Fifty pl of 3 M sodium acetate and 1 ml of ice cold 100% ethanol were added to this solution to precipitate the DNA, which was then pelleted down after being stored for 2 hours at -20°C. Washing with 70% ethanol was carried out twice to desalt the DNA pellet, which was then dissolved in 200 pl of TE buffer with 10 pg of RNase A being added. After incubation at 37°C overnight, the DNA solution was extracted with phenol/chloroform/ isoamyl alcohol (24:24:1) once, and DNA was precipitated with three volumes of 100% ethanol. The DNA pellet was dried using SpeedVac and re-dissolved in 500 pl TE buffer for quantification and further applications. Polymerase Chain Reaction (PCR) was employed to screen plants for transgene integration. PCR primers were designed using Oligom‘4.0 software (Molecular Biology Insights, 79 Inc.) on a Macintosh computer and were synthesized by Michigan State University Macromolecular Facility using ABI 3948 System (PE Applied Biosystem). Sequences of these primer pairs are: [pHSZ] upper primer (MB56) 5’- TCTACTACCCAGCGAAAC-3’ and lower primer (M855) 5’- CCTTTGTTCTTATTCCATT-B’, which prime the synthesis of the fragment of 1136 base pairs between position 30 and 1166 in the H82 coding sequence (Hajela et al., 1993, unpublished, Genbank accession: L22032); [bar gene] upper primer (MB76) 5’-TACCGAGCCGCAGGAA-3’ and lower primer (MB77) 5’- TGTAGAGCGTGGAGCC-3’, which prime the synthesis of the fragment of 189 base pairs between position 270 and 466 in bar gene (Thompson et al., 1987, Genbank accession X05822); [mtlD] upper primer (MB 59) 5’-AATATCGGTCGTGGCT-3’ and lower primer (MB60) 5’-CTTTGTCAGCGATCAGT-3’, which prime the synthesis of the fragment of 1029 base pairs between position 182 and 1211 in the mtlD sequence (Jiang et al., 1990, Genbank accession X51359). DNA amplifications were conducted in a 96—well Thermal Cycler (Perkin-Elmer, Norwalk, CT). The thermal cycles are: for pHSZ gene, 94°C 4 min.(pre-denaturing), 35 cycles of 94°C 30 sec.--- 52°C 30 sec.--- 72°C, and 72°C 5 min. (post-extension); for bar gene, 94°C 4 min. (pre-denaturing), 35 cycles of 94°C 15 sec.--- 60°C 30 sec.--- 72°C, 72°C 5 min.(post-extension); 8O for mtlD, 94°C 4 min. (pre-denaturing), 35 cycles of 94°C 30 sec.--- 54°C 30 sec.--— 72°C, 72°C 5 min. (post-extension). AmpliTaq (Perkin Elmer) polymerase was used for PCR. DNA slot blotting was carried out for estimating relative transgene copy number using a MinifoldC>III slot- blot system (Schleicher & Schuell Inc., Keene, NH). A piece of 20 X SSPE buffer (3.6 M NaCl, 0.4 M NaHPO4, 0.04 M EDTA, pH 7.7) pre—wetted HybondT"-N+ nylon membrane (Amersham Corp., Arlington Heights, IL) was used to bind DNA. Five micrograms of DNA from non-transgenic control and each of the transgenic lines were loaded in each slot. Plasmid standards equivalent to 1, 2, 4, and 8 copies of the transgene per genome were loaded. The hybridization procedure for DNA slot blot was the same as that described for Southern blot below. The bands were read and analyzed using Computing DensitometerTM and ImageQuantTM 3.5 (Molecular Dynamics, Inc., Sunnyvale, CA). In Southern blot analysis, restriction digestion of the plant and plasmid DNA was carried out using restriction enzymes from Boehringer Mannheim Corp. (Indianapolis, IN). Twenty pg of plant DNA from each sample were digested overnight in microcentrifuge tubes with 100 units of one or two restriction enzymes in a total volume of 500 pl. After being dried down to 200 pl in SpeedVac, each digestion was 81 added three volumes of 100% ethanol to precipitate DNA pellet, which was then desalted by two washes of 70% ethanol and subsequently dried in SpeedVac. The DNA pellets in each tube were dissolved in 20 pl of 1 X DNA loading buffer and loaded on an 0.8% agarose gel. Electrophoresis was carried out at 20 volts for 16 hours. After completion, the gel was soaked in 0.2 N HCl for 8 minutes for depurination and then rinsed briefly with distilled water. The gel was then denatured with two changes of the buffer containing 1.5 M NaCl and 0.5 M NaOH for 15 minutes each time, and neutralized in a buffer containing 1.5 M NaCl and 1 M Tris-HCl (pH 7.5) for 30 minutes. The treated DNA in the agarose gel was transferred onto a pieces of pre-wetted Hybondm-N+ nylon membrane (Amersham Corp., Arlington Heights, IL) by capillary flow of 10 X SSPE buffer (1.8 M NaCl, 0.2 M NaHPO4, 0.02 M EDTA, pH 7.7) for about 18 hours. When the transfer was complete, the nylon membrane was peeled from the gel and soaked into 5 X SSPE buffer for 5 minutes to remove any pieces of agarose sticking to it. The nylon membrane was then baked in a vacuumed oven for 2 hours at 80°C. DNA fragments to be used as probes were fractionated from restriction digestions of plasmid DNA by agarose gel electrophoresis. Gel bands containing the fragments of 82 interest were visualized under UV illumination and excised from the gel, and then were treated for elution and purification using GENECLEANC>II kit (Bio 101, Inc., Vista, CA). Probe DNA was labeled with deoxycytidine 5’- triphosphate (dCTP) [a- 32P] (specific activity 3000 Ci/mmol) using'TJQuickPrinmJ”IKit (Pharmacia Biotech, Inc., Piscataway, NJ) and the protocol provided by the company. The free labels 32P were then removed from the probe solutions after passing through gel columns made of SephadexC>G-5O (Pharmacia Biotech, Inc.). The nylon membrane carrying the DNA blot was first wetted with distilled water and incubated at 68°C for 1 hour with the hybridization buffer (7% SDS, 0.25 M Nafimh, pH 7.4) in a lidded plastic container without the probe. The membrane was rolled into the hybridization tube (Robbins Scientific Corp., Sunnyvale, CA) containing a full load of the pre- heated hybridization buffer (68°C). The air bubbles trapped between the membrane and inside of the hybridization tube were smoothed away with a glass rod and all but 3-5 ml of the hybridization buffer was poured out of the tube. The labeled probe was then add to the buffer in the tube, which was then loaded onto the rotator in a hybridization incubator (Robbins Scientific Corp., Sunnyvale, CA) and rotated & incubated at 68°C overnight. Three washes (15 83 minutes each) were carried out at 68°C with the wash buffer (0.1 X SSPE and 0.1% SDS) in the glass tube. The washed membrane was then wrapped with plastic film and placed in a cassette with a piece of 8 1/2 X 11 X-OMATW film (Eastman Kodak Company, Rochester, NY) for radioautography. RNA Isolation and Analyses A modified protocol following Extract-A—PlantTM RNA Isolation Kit (Clontech Laboratories, Inc., Palo Alto, CA) was used to isolate total RNA from creeping bentgrass. Four milliliters (4 ml) of extraction buffer (100 mM LiCl, 100 mM Tris-HCl pH 8.0, 10 mM EDTA, 1% SDS) and an equal volume of water-saturated phenol were dispensed into a 15 ml sterile polypropylene centrifuge tube (Corning, Inc., Corning, NY) and heated to 80°C in a water bath. A chilled mortar and pestle that had been baked at 180°C overnight were used to grind 2 g of young leaf tissue of the plants in liquid nitrogen. The ground leaf powder was transferred to the centrifuge tube containing the preheated phenol and extraction buffer. Vortexing for 30 to 60 sec. was carried out to homogenize the liquid-powder mixture and then 2 ml of chloroform : isoamyl alcohol (24:1) was added, followed by vortexing for another 30 to 60 sec. The extraction mixture was centrifuged for 10 minutes at 2275 x g (4235 84 rpm) in a IEC 6 x 15 ml centrifuge (International Equipment Company, Needhamhts, MA) to separate the organic phase containing plant tissue debris from the upper aqueous phase, which was then transferred to a 15 ml Corex® 11 glass centrifuge tube (Corning Inc., Corning, NY) that had been baked at 180°C overnight. One volume (4 ml) of 4M LiCl was added to and mixed with the aqueous RNA-containing solution in this Corex® II tube, which was then placed in an upright position in a -80°C freezer for 1 hour before being centrifuged at 10,000 g for 20 minutes. The RNA pellet was washed twice to remove the trace LiCl with 70% ethanol, then air dried, and finally dissolved in 1 ml diethyl pyrocarbonate (DEPC) (Sigma Chemical Co., St. Louis, MO) treated HAD. To prevent cross contamination, all isolation and purification steps for RNA samples to be used for RT-PCR were performed in microcentrifuge tubes. mRNA species were purified using PolyATtract® mRNA Isolation System IV (Promega, Madison, WI). RT-PCR for each sample was performed in two separate tubes. In reverse transcription, 4 pl total RNA (1 pg/pl), 4 pl lower primer (5 pM), and llLlng were mixed up in a microcentrifuge tube, heated at 65°C for 5 minutes, and then chilled on ice. Then, the following components were added: 4 pl M-MLV buffer (5 X), 4 pl dNTP (10 mM), 2 pl DTT, and l 85 pl M-MLV reverse transcriptase (RT) (200 unites/pl) (Gibco BRL, Rockville, MD). This reaction mix was incubated at 42°C for 2 hour, and then heated at 95°C for 5 minutes to inactivate the RT enzyme. From the RT reactions, 2 pl from each sample was used to perform regular PCR in the 96-well thermal cycler (Perkin-Elmer, Norwalk, CT). Protein Isolation and Analyses About 0.2 grams of fresh leaf tissue was chilled in liquid nitrogen and ground into fine powder in a microcentrifuge tube using a disposable pestle. 0.4 ml of protein extraction buffer (100 mM sodium acetate, adjusted to pH 5.0 using HCl) was dispensed into each sample, and then vortexed for 2 minutes. The samples were then placed on ice and incubated for 15 minutes before centrifugation at 2,000 g for 5 minutes. The supernatant from each tube was transferred to a clean tube and stored at -80°C. The concentration of total protein in each sample was measured using the Bradford method with Bio-Rad protein assay reagent (Bio-Rad, Hercules, CA). One hundred pg of total protein from each sample was mixed with 5 X protein loading buffer (250 Mm Tris-HCl, pH 6.8, 500 mM dithiothreitol, 10% SDS, 0.5% bromophenol blue, 50% glycerol) and heated at 95°C for 10 minutes. The cooled 86 samples were loaded onto 10% SDS polyacrylamide gel (resolving gel), and run in Mighty Small II vertical gel apparatus (Hoefer Scientific, San Francisco, CA) under 100 volts until the tracking dye reach the bottom of the gel. The gel was then electro-blotted onto a piece of BA-S NC nitrocellulose membrane (Schleicher & Schuell, Keene, NH) under 20 volts at 4°C overnight with stirring and cooling water circulation. The blotted membrane was then incubated with gentle swirling for 1 hour at 6°C in 1 X TBST (150 mM NaCl, 10 mM Tris-HCl pH 8.0, 0.05% Tween 20) containing 5% (w/v) nonfat milk and 0.02% sodium azide. The primary antiserum raised against a H82 peptide-containing glutathione S-transferase (GST) fusion protein (Pharmacia, Biotech Inc., Piscataway, NJ) from a rabbit was then added to the buffer to a final dilution 1:200 and incubated with the membrane for an additional 5 hours. The membrane was then incubated with three changes of 1 X TBST buffer for a total of 45 minutes. The secondary antiserum (alkaline phosphatase conjugated anti-rabbit IgG, Sigma Chemical Co., St. Louis, MO) was dispensed into 1 X TBST buffer to the final dilution of 1:7500 and incubated with the membrane at 6°C for 2 hours with gentle shaking. Three washes of the membrane were carried out using 1 X TBST similar to previous ones. The membrane was put in alkaline phosphatase 87 buffer (100 mM Tris-base, 100 mM NaCl, 5 mM MgC12, pH 9.8) containing 0.33 mg/ml of nitro blue tetrazolium (NBT) and 0.16 mg/ml of 5-bromo-4-chloro-3-indolyl phosphate (BCIP) for color development. 88 RESULTS Regeneration of Herbicide Resistance Creeping Bentgrass Eight gene delivery experiments were carried out using BiolisticC>Particle Delivery System. A total of 260 callus and suspension preparations were bombarded with tungsten particles coated with plasmids pKYLX71-H82 and pJSlOl. Callus cultures from solid medium and suspensions were used to prepare for the bombardment. Preparations that had been bombarded were immediately transferred to plates with fresh callus induction medium containing no selection agents. Four to five days after gene delivery, callus induction medium containing the selection agent bialaphos was used for the first two subcultures. To ensure that the selection was carried out on every cell, larger pieces of callus were divided into smaller ones of about 1 mm in diameter. Selection in the callus was allowed for 12 weeks in callus selection medium with first two subcultures using medium containing 5 mg/L and the third subculture using medium containing 10 mg/L of bialaphos in the dark. The surviving calli (light colored, and vigorously growing) were transferred onto plant regeneration containing 15 mg/L bialaphos and placed under light for regeneration. A total of 47 of 109 bialaphos-resistant callus lines were able to 89 Table 2. Bialaphos Selection and Plant Regeneration after Bombardment No. of surviving Callus clumps and plantlets on Number of Exp. Preparations bialaphos medium resistant I Bombarded (# Resistant Viable clones per 0f plates) calli plants plate Week 4 Week 8 Week 12 Week 16 Callus 0r 5 mg/L 5 mg/L 10 mg/L 15 mg/L Suspension 1 Callus 10 10 2 1 1 0.10 Suspension 10 3 0 0 0 0.00 2 Callus 10 12 7 4 2 0.20 Suspension 10 19 7 6 0 0.00 3 Callus 10 l9 l8 3 3 0.30 Suspension 10 26 24 10 7 0.70 4 Callus 10 12 9 5 3 0.30 Suspension 10 9 5 5 4 0.40 5 Callus 12 15 12 7 5 0.41 Suspension 12 22 17 4 3 0.25 6 Callus 12 21 20 20 7 0.58 Suspension 12 16 15 15 4 0.33 7 Callus 12 15 10 9 2 0.17 Suspension 12 15 13 12 3 0.25 8 Callus 12 11 9 3 3 0.25 Suspension 12 7 7 5 0 0.00 Sub— toLa1 Callus prep. 115 87 52 26 Sub— total Suspension 117 88 57 21 prep. Average Total 176 232 175 109 47 0.26 147/176) 90 regenerate into viable plants (Table 2), among which 8 derived from Penncross and 38 from Putter. All these clones trace back to independent callus lines. A significant number of callus lines were not able to give rise to viable green plantlets on the selection medium. Some of them remained as callus structures, some produced albino plantlets that eventually died, others produced green plantlets but were later eliminated by selection at the plantlet stage. All but one of the plantlets that survived the selection at the highest concentration of bialaphos (15 mg/L) also survived subsequent sprays in the greenhouse using 1.2% Ignite“fl Some morphological abnormalities were observed from both Penncross- and Putter- derived bialaphos-resistant lines. There were 5 clones grew slower by two to three fold than the rest of the transgenic lines and the seed germinated plants, which has an average leaf length growth rate of about 4 cm/week at 4.4 g/HE/week of N2 (1/4 pound per thousand square feet). Two of these clones also tended to be “shooty” by having long shoot internodes and relatively short and coarse leaf blades, while the other transgenic lines and nontransgenic parents were more prostate with shorter internodes and slender leaves. 91 Transformation and cotransformation as confirmed by DNA analyses Polymerase Chain Reaction (PCR) was used to evaluate transgene integration in the originally putative transgenic lines. Fragments of 189 bp (bar) (Figure 2), 1166 bp (H82 gene)(Figure 3), and 1029 bp (mtlD gene)(Figure 4) were amplified using DNA isolated from leaf tissues of the bialaphos resistant plants and non-transgenic control plants. Two hundred ng of template DNA was using for each. reaction. PCR results for bar gene integration were consistent with the resistance of the plants to Ignite herbicide sprays, which eliminated one escape from bialaphos selection. DNA slot blots provided another means for detecting the presence of specific sequences in DNA samples. Five micrograms of DNA from each of 35 individual bialaphos resistance plants were loaded in each slot. The results are shown in Figure 5. The relative copy numbers of gene insertions were estimated based on the density of each slot in comparison to plasmid controls. Copy numbers of the three transgenes differed significantly among these transgenic lines from zero to more than ten copies. 92 12 3 4 5 6 7 8 91.0111213141516171819202122 189 bp 12345678910 Figure 2. PCR detecting bar gene in creeping bentgrass transgenic plants. Upper Lane 1: molecular weight markers. Lane 2, 3,and 4: non-transgenic plants. Lane 5 through 22: bialaphos resistant plants. Lower Lane 1 and 10: molecular weight markers. Lane 2 through 8: bialaphos resistant plants. Lane 9: plasmid pJSlOl positive control. A fragment of 186 bp was amplified as indicated by the arrow. 93 ‘— 1.1kb 1 2 3 4 5 6 7 8 9 10111213141516171819202122 Figure 3. PCR detecting H82 gene in creeping bentgrass transgenic plant. Upper and Lower Lane 1: Molecular weight markers. Lane 2: non-transgenic plant. Lane 3. Plasmid pKYLX71—H82 positive control. Upper Lanes 4 through 22 A 1.0 kb fragment within H82 coding sequence was amplified as indicated by the arrow. 94 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 ‘— 1.0 kb ‘— 1.0 kb 1 2 3 4 5 6 7 8 9 10 ll 12 13 14 15 16 17 18 19 20 21 22 Figure 4. PCR detecting mtlD gene in creeping bentgrass transgenic plants. Lane 1: Molecular weight marker. Lane 2: Non—transgenic plant. Lane 3: plasmid pJSlOl positive control. Upper Lanes 4 through 22: bialaphos resistant plants; Lower Lanes 4 through 21: bialaphos resistant plants. A 1.0 fragment was amplified as indicated by the arrow. 95 2122 232425$2728293031323334353637383940 1234567891011121314151617181920 21222324252627 28293031323334353637383940 1234567891011121314151617181920 21222324 252627282930 31 3233343536373839 40 l2341567891011121314151617181920 Figure 5. Slot blot analyses. Slot 1: Non-transgenic plants. Slot 2 through 35: independent transgenic lines. The same samples were loaded in slots with the same number throughout A, B, and C sections. Slot 36: blank control. Slot 37 through 40: plasmid controls (in the order of 1x, 2x, 4x, 8x dosage). A. H32 detection. B. mtlD probe detection. C. bar probe detection. 96 Southern blot analyses were performed to further examine the H82 and mtlD gene insertions in the genomic DNA of these transgenic plants. Twenty micrograms of three DNA preparations from each sample, undigested, single cut, and double digested DNA by restriction enzymes, were loaded onto parallel to the electrophoresis gel. DNA from non- transformed plants and plasmids were used as the negative and positive controls, respectively. Based on the genomic size for creeping bentgrass species (Bennett and Leitch, 1995) and the sizes of the plasmids, pJSlOl (8.5 kb) and pKYLX71-H82 (12.0 kb), the amount of plasmid DNA for each' positive control lane was calculated to be equivalent to 20 pg of creeping bentgrass genomic DNA that contains one copy, three copies, and five copies of mtlD or H82 gene. PCR and Southern blot analyses confirmed all but one bialaphos resistant lines to have mtlD gene insertions (Figure 6). Eleven of the 18 slot blot positive lines had been confirmed to have H82 gene insertion(s) in their genomic DNA by PCR and Southern blot analyses. The unique banding pattern in lanes with single digested DNA indicated their status as independent transgenic lines (Figure 7). The cotransformation frequency, which was defined as the frequency with which cells transformed with bar 97 contained a H52 or mtlD gene, was calculated. Based on data generated from DNA analyses, the cotransformation frequency of bar (selectable marker) and its linked mtlD gene (unselected gene) was 100% while the cotransformation frequency of bar (selectable marker) and its unlinked H82 gene (nonselectable gene) accounted for 23.9% of the total bar-containing lines (Table 3). Significant correlation was observed between the hybridization density counts of the co—inserted bar and mtlD gene using densitometer readings of the slot blot radioautographic images. The correlation coefficient of bar and mtlD genes densitometer readings was estimated as r = 0.7913 and tested significant (a < 0.05) using Weir’s two- sided T-test formula(r > 2/7‘3’ ). Surprisingly, correlation also existed between bar and the H82 gene based on correlation coefficients, which r = 0.8587 and the T-test using the same formula and standard (Figure 8). RNA Analyses RT-PCR analyses were conducted as a sensitive tool to screen the transgenic plants for transgene expressions at the mRNA level. The same pair of primers for PCR of the H82 gene was used for RT—PCR amplifications. The results are shown on Figure 9. 98 Table 3. resistant transgenic lines Gene integration and expression in bialaphos Gene integration Gene expression PCR Slot Souther RT- Northern Western blot n blot PCR blot blot bar 46/47 33/34 ---------- 10/11 ----- mtlD 46/47 33/34 39/40 ----- 10/11 ----- H82 11/47 18/34 11/20 11/11 6/11 4/11 bar-mtlD 100% bar-HSZ 23.9% Cotransformation frequency (46/46) (ll/46) (based on PCR): 99 __J1____ _NI_ _J&_ _JL_ __C_ _JL_ 1 2 3 1 2 3 l 2 3 1 2 3 1 2 3 l 2 3 L5 kb ‘— __E_.._JNI _JL_ _E__ _IL_. _IL_. __1_. _l__ 1 2 3 1 2 3 1 2 3 l 2 3 1 2 3 1 2 3 1 2 3 1 2 3 Figure 6. Southern blot analyses of transgenic creeping bentgrass for mtlD gene insertions. P: DNA of plasmid pJSlOl serving as positive controls. NT: DNA of nontransgenic creeping bentgrass plants serving as negative controls. A, B, C, D, E, F, G, H, I, and J represent different transgenic lines. Lane 1: undigested genomic DNA; Lane 2: Hind III digested genomic DNA; Lane 3: Hind III and Bam HI digested genomic DNA. A 1.5 kb fragment of pJSlOl from HindIII/BamHI digestion was used as the probe. This mtlD coding region was released by restriction digestion as indicated by the arrow. __E__ _JEI. _JL__ __IL_ __IL_. __IL__ __E___ _ 1.2.3 1 2 3 1 2 3 1 2 3 1 2 3 1 2 3 1 2 3 .9 X s N W O" _JL_ NT F G 11 I J 1 2 3 1 2 3 1 2 3 1 2 3 1 2 3 1 2 3 1 2 3 .- "my ' . - I .. " . a "II 0 L2kb Figure 7. Southern blot analyses of transgenic creeping bentgrass for H82 gene insertions. P: DNA of plasmid pKYLX71-HSZ serving as positive controls. NT: DNA of nontransgenic creeping bentgrass plants serving as negative controls. A, B, C, D, E, F, G, H, I, and J represent different transgenic lines. Lane 1: undigested genomic DNA; Lane 2: EcoRI digested genomic DNA; Lane 3: PstI digested genomic DNA. A 1.2 kb fragment of pKYLX71-HSZ from PstI digestion was used as the probe. This H82 coding region was released by restriction digestion as indicated by the arrow. 101 Densitometer Reading of Slot Blot . um fl 5 mm a mm .2 H 5 25° , . amtID g 200 . " lbar «9 150 ' “$2 -- ‘wo x 2 8X 31 50 8X 3 DNA sample 2.839 59.75 0.5 23.67 3.365 12.61 149 40.93 87.3 117.9 34.16 94.92 66.48 35.97 94.58 0.5 157.6 15.05 149.8 94.06 19.04 56.46 31.3 6.584 178.3 9.878 60.2 7.476 25.42 52.72 3.445 4.033 7.194 0.5 42.9 27.81 DNA sample Figure 8. Correlation of insertion dosage among the three transgenes for 35 clones of transgenic creeping bentgrass. The numbers in the table are the hybridization density counts from each sample. “8 X” indicates the levels of hybridization density count equivalent to 8 copies of gene insertion in the transgenic plants. 102 Northern analyses were used to detect the actual levels of mRNA products from gene transcriptions. As expected, all transgenic lines showed detectable levels of bar mRNA transcripts using 20 pg of total RNA (Figure 10). There was no apparent correlation between mRNA levels of bar gene and its respective copy number. For the H82 gene, mRNA signals could be detected from only two lines, 8151 and 9606 (Figure 10. Lane E and Lane F), by using 20 pg of total RNA, while using enriched mRNA samples enabled northern blot to detect an additional four transgenic lines containing the H82 gene insertions (Figure 10. Lane A:91012, Lane B: 8157, Lane C: 711, Lane D: 9603, Lane E: 8151, Lane F: 9606). Immunoblot (Western Blot) Analyse Immunoblots were performed to confirm the chitinase synthesis as the result of the H82 gene expression at the protein level. Using rabbit antiserum raised from a rabbit against a GST—chitinase fusion protein, low levels of chitinase protein were detected from four of the six transgenic lines expressing the H82 mRNA (Figure 11. Samples in Lane 1, 2, 3, 4, 5, 6, and 7 correspond to Lane A, B, C, D, E, and F in Figure 10). 103 Assays of Brown Patch Resistance Two of creeping bentgrass clones expressing the H82 gene, 711 and 9603, from this study were found to be three and one—fold more resistant, respectively, to infections of brown patch disease (Rhizoctonia solani) in growth chamber and greenhouse settings (the testing work was conducted by our research collaborators in Dr. Joseph Vargas’s lab, and the results will be published in Journal of the American Chemical Society). 104 4— 1.1kb WA182C3D4E5F6G7H8I9P1P2 Figure 9. RT-PCR detecting H82 transcripts. Lane mw: molecular weight markers. Lane A: RT-PCR of non-transgenic plant. Lane 1: PCR of non—transgenic plant. Lane A, B, C, D, E, F, G, H, and I: RT-PCR of transgenic plants. Lane 1, 2, 3, 4, 5, 6, 7, 8, and 9: PCR of the transgenic plants. P1 and P2: PCR of plasmid pKYLX71-HSZ. A fragment of 1.1 kb was amplified as indicated by the arrow. 105 NT A B C D E F G H I J K NT A B C D E F G H I J K TC T NT E F NT A B C D E F Figure 10. Northern blot analyses of respective creeping bentgrass transgenic plants. Top: bar gene transcripts. Middle: mtlD gene transcripts. Bottom: H52 gene transcripts. Lane NT: non-transformed control. Lane A through K: independent transgenic lines. Lane TC: non- transformed tobacco plants. Lane T: transgenic tobacco with H82 gene. DNA probes for all three genes were the same used in Southern blots. 106 Figure 11. Western blot analyses of the transgenic creeping bentgrass expressing the HS2 gene. Lane MW: molecular weight marker. Lane 1: non-transgenic plant. Lane 2 through 7: independent transgenic lines. Lane 8 and 9: transgenic tobacco. The arrow indicates the protein band of chitinase. 107 DISCUSSION The results demonstrated the effectiveness of using a selectable marker gene to facilitate selection of creeping bentgrass transformed with a non-selectable gene after they were co-introduced into target cells. Cotransformation has been used more and more frequently in delivering nonselective genes into plants. Komari et al. (1996) used transformed rice and tobacco plants using 'super—binary' vectors that carried two separate T-DNAs via Agrobacterium tumeifaciens, each of which contained a drug resistance selectable marker gene and a GUS gene. They achieved cotransformation frequencies up to 47% and obtained marker- free plants from the segregating progeny. Buck et al. (1998) also achieved similar cotransformation of two unlinked genes, and they suggested that the occurrence of marker-free shoots and progeny might result from the expression of unstable, transiently expressed selectable marker gene in the cells, which enabled them to survive the selection. Cotransformation also proved effective in microprojectile-mediated gene transfer for maize (Gordon- Kamm et al., 1990) and sugarcane (Bower et al., 1996). The cotransformation frequency (23.9%) for unlinked bar and the H82 gene from our study was similar to that reported 108 previously for other species (Gordon-Kamm et al., 1990; Christou and Swain, 1990; Damm et al., 1989). These reports also indicate that linkage of the selected and unselected genes on a single plasmid has led to a significantly higher cotransformation frequencies than was reported with unlinked genes. On the other hand, while the frequency of cotransformation between unlinked bar and the H52 gene in this research was substantially lower than the linked cotransformation, the significant correlation between copy numbers of bar and the H52 insertions may suggest the occurrence of the gene integration events in some proximity. The only drawback of unlinked cotransformation, its lower frequency, could be over-compensated by its convenience value because the unlinked cotransformation precludes the necessity of constructing a specific plasmid harboring both the selectable marker and each of the economically desirable genes. Another advantage of cotransformation is the ease of obtaining selectable marker-free plants from the segregating progeny (Komari et al., 1996; Buck et al., 1998). Another important finding from this study was the fact that expression level of the H52 gene was low or not detected in some transgenic lines containing this gene. 109 There are two potentially plausible explanations for this phenomenon. Que and Jorgensen (1998) suggested that transgenes might be involved in two types of homology- dependent gene silencing (HdGS) mechanisms. One requires promoter homology, the other requires transcript homology. Gene silencing mediated by 355 promoter was reported by Park et al. (1996) in transgenic tobacco. In their study, the expression of hygromycin phosphotransferase (hpt) gene driven by a 355 promoter was inhibited because of the presence of another 355 promoter at locus 271. This type of gene silencing occurred at the transcriptional level and was accompanied by a reduction in the steady-state levels of hpt transcript. Because the 355 promoter was used in the creeping bentgrass research here to initiate gene transcription for both bar and the H52 gene coding sequences, it is possible that this kind of trans acting inhibition happened to plants carrying both bar and the H52 gene insertions. Methylation of the promoter sequence was determined as the mechanism for gene silencing of this type (Park et al., 1996; Kumpatla et al.,1997; Que and Jorgensen, 1998). Genomic sequencing of the H52 insertions may be able to shade light on the underlying mechanisms of this phenomenon observed in my study. Analyses of the progeny for the H52 gene expression will also help us to 110 understand the real cause, because of the heritable nature of DNA methylation. One other way to study the 355 promoter mediated gene silencing in my case is to substitute the H52 gene in plasmid pKYLX71-H52 with a selectable marker gene and co- transfer it with pJSlOl (where the bar gene is housed) into creeping bentgrass plants. Selections using bar and the new selectable marker should be carried out separately. If this hypothesis turns out to be valid, we should be able to see the inhibited expression of bar gene in plants selected' with the new marker, and vice versa. The other possible explanation for the low level for transcription of the H52 gene is the co—suppression of gene or silencing effect between native chitinase genes and their products and the H52 gene and it products. This possibility exists because chitinase genes have been detected or isolated from other turfgrass species related to creeping bentgrass (Roberts et al., 1992; Anderson et al., 1997; Reyes, 1999). Some morphological variations had been observed among the transgenic plants. It was not surprising, considering the heterogeneous genetic background of the plant material used and the likelihood of somaclonal variation incurred from in vitro manipulations at the callus phase. 111 This study has expanded the application of the transformation system developed by Zhong et al. (1993) and the bar gene selection scheme formulated by Liu (1996), which previously proved to be effective in linked cotransformation, to unlinked cotransformation of creeping bentgrass. Two of the H52-expressing lines, 711 and 9603, exhibited enhanced resistance to infection by the brown patch pathogen (Rhizoctonia solani). This result suggests that the synthesized transgenic chitinase protein might“ have conferred antifungal activity to these two transgenic lines. However, it is not clear why the other two H52— expressing lines, 8151 and 9606, did not show the similar resistance to the brown patch pathogen. 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