allelic composition and genetic background effects on transgene expression and inheritance in white...

Molecular Breeding4: 479–490, 1998.© 1998Kluwer Academic Publishers. Printed in the Netherlands.

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Allelic composition and genetic background effects on transgeneexpression and inheritance in white clover

Alicia Scott1,∗, Derek Woodfield2 & Derek W.R. White1AgResearch, Private Bag 11008, Palmerston North, New Zealand1Plant Molecular Genetics Laboratory and2Plant Improvement, Grasslands Division, AgResearch, Private Bag11008, Palmerston North, New Zealand (∗author for correspondence; e-mail:[email protected])

Received 1 December 1997; accepted in revised form 27 May 1998

Key words:breeding strategy, gene dosage, genetic background, inheritance, transgene expression, transgenicwhite clover

Abstract

Allelic composition and genetic background effects on GUS expression and inheritance using a chimeric(cauliflower mosaic virus 35Sp:uidA) transgene were investigated in white clover as a prelude to transgenic cultivardevelopment. Stable expression and Mendelian inheritance of theuidA transgene was observed over two genera-tions when theuidA transgene was maintained in a heterozygous state. Transgenic backcross progeny (BC1) wereintercrossed to produce segregating F2 populations. GUS-positive F2 plants were test-crossed with a non-transgeniccontrol plant to determine whether individuals were heterozygous or homozygous for the transgene. Both expectedand distorted segregation ratios were observed. Distortion of the segregation ratio was not caused by transgeneinactivation or rearrangement, but was influenced by genetic background. BC1, BC2 and F2 populations werefound to have similar levels ofuidA gene expression. Quantification of GUS expression from progeny of highand low GUS expressing plants indicate that it is possible to alter transgene expression through selection. Nodifference was found between the level of expression for F2 plants homozygous or heterozygous for the transgene.These results indicate that F2 plants, homozygous for a transgene, might be used to develop a transgenic cultivar.However, progeny testing to determine the influence of genetic background is a prerequisite to such a development.

Introduction

White clover is the most important pasture legume inmany temperate regions of the world and has beensignificantly improved by traditional breeding meth-ods in the past 60 years [33]. Recombinant DNAtechniques can be utilised to develop transgenic whiteclover cultivars with traits previously not found withinthe natural genetic variation of this species. Initially,white clover was a difficult species to transform, inthat transformed plants could only be produced froma single genotype [30]. Now, a wide range of geno-types are routinely transformed [28, 19] and the firstfield trial of transgenic white clover plants has beenundertaken [10]. However, before a transgenic whiteclover cultivar can be released, stable inheritance andexpression of the transgene must be demonstrated.

Duan et al. [9] demonstrated in transgenic riceplants that an introduced potato proteinase inhibitor IIgene was stably inherited and expressed for more thanfour generations. Mülleret al. [22] also have foundthat transgenes are transmitted to progeny with a highdegree of meiotic stability. However, others have re-ported instability of transgene inheritance, includingtransgene loss or rearrangement, whether plants weretransformed by PEG-mediated direct gene transfer [5],biolistic bombardment [27] orAgrobacteriummedi-ation [6]. Srivastavaet al. [27] revealed a potentialproblem in the breeding of transgenic cultivars sinceinstability of the transgene was not apparent until thesecond or third generation. In this instance the trans-gene segregated as expected in the first generation butsubsequently lost expression and was eliminated fromthird-generation plants.

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Unlike species such asNicotiana, Petunia andArabidopsisused as model systems in transforma-tion studies, white clover is an outcrossing speciesand predominantly self-incompatible. Populations andcultivars are therefore a heterogeneous mixture ofheterozygous individuals, resulting in high levels ofgenetic variation both within and among populations[32]. This variability contributes to the broad environ-mental adaptation and phenotypic plasticity of whiteclover [34]. In developing a transgenic white clovercultivar, it is desirable that this genetic heterogeneitybe maintained.

Ideally, an introduced gene should be expressed inevery genotype of a transgenic cultivar. This can beachieved by having the transgene in a homozygousstate, therefore ensuring all progeny of the next gen-eration will inherit the transgenic trait. Since whiteclover is predominantly self-incompatible we devel-oped a strategy to cross a single primary transgenicplant to a range of plants to produce backcross popu-lations (BC1), then intercross transgenic BC1 plants toproduce F2 populations. An F2 population will have ahighly heterogeneous genetic background and a vari-able allelic copy number (0 to 2) for the transgene.Therefore, it is essential to establish the effects ofboth allelic composition at individual loci and differ-ing genetic backgrounds on transgene expression andinheritance. Within an F2 population, plants homozy-gous for a transgene can be identified by test-crossing.By inter-crossing homozygous F2 plants between thedifferent F2 populations it is possible to preserve thegenetic heterogeneity of the white clover cultivar butfix the transgene so that it is expressed by everyindividual within the cultivar.

In this study, we tested the stability and inheritanceof the same transgene in different white clover geneticbackgrounds as a prelude to transgenic white clovercultivar development. Inheritance and expression ofthe transgene was examined in the heterozygous andhomozygous state. The effect of selection on trans-gene expression was also investigated. We used aprimary transgenic plant (GH7) we had previously de-scribed [28] since this plant contained a single T-DNAinsert, exhibited strongβ-glucuronidase (GUS) activ-ity, and backcross progeny from this plant segregatedfor GUS activity according to Mendelian inheritance.The T-DNA contained aβ-glucuronidase (uidA) re-porter gene and a neomycin phosphotransferase II(nptII gene) to confer antibiotic resistance. TheuidAgene was under the control of a truncated (−288 to+2) CaMV–35S promoter, whilst the selectablenptII

gene was fused to a full-length CaMV–35S promoter.Both genes contained 3′ octopine synthase (OCS)polyadenylation regions.

Materials and methods

BC1, and BC2, progeny

The primary transgenic plant (GH7) was crossed witha non-transgenic control (GH9) to generate a back-cross (BC1) population (Figure 1). Both parentalplants were derived from cv. Grasslands Tahora.Reciprocal crosses were made on emasculated plantshand pollinated in a glasshouse.

The four highest expressing GUS plants (GH646,GH652, GH694 and GH708) were identified withinthe BC1 population and outcrossed to a non-transgeniccontrol plant (C6525/13 cultivar Grasslands Sustain)to produce a BC2H population. An unrelated controlplant was chosen to avoid any possibility of inbreed-ing depression influencing expression and inheritanceof the transgene. Similarly the four lowest-expressingplants (GH633, GH637, GH644 and GH698) wereused to generate a BC2L population. Non-reciprocalhand pollinations were performed on unemasculatedplants with the Grasslands Sustain plant used as apollen donor. All maternal plants were self-pollinatedas a check for possible self-compatibility.

F2 allelic composition determination

Five of the BC1 progeny that exhibited high GUS ac-tivity were intercrossed to produce a segregating F2population. GUS-positive F2 plants were test-crossedwith a non-transgenic control plant (I2/6 or I2/18 cv.Crau) to determine whether the plant was heterozy-gous or homozygous for the transgene. The F2 plantswere used as female parents in the test-crosses. Twentyseedlings from each GUS-positive F2 plant were histo-chemically assayed for GUS activity.In vitro seedlingswere tested after 1 and 4 weeks in culture. F2 plantswere classed as homozygous for theuidA transgene ifall 20 progeny either exhibited GUS activity or wereshown by PCR techniques to have inherited theuidAtransgene. For segregating populations, two GUS-positive and three GUS-negative plantlets were testedfor the presence of theuidA transgene using PCR.

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Figure 1. Crossing scheme used to generate the BC1, BC2, F2 and test cross progeny. AA denotes a diploid chromosome set whilst an∗ thepresence of theuidA transgene within a chromosome. For populations that segregated for the presence of the transgene, only GUS-positiveindividuals (underlined) were crossed to produce the next population.

Tissue culture

All seed was germinatedin vitro. Seed was germi-nated and plantlets maintained at 26◦C under a 16 hphotoperiod of 55 to 85µE m−2s−1 of light. Seedwas surface- sterilised with acidified HgCl2 [31]. Aftersterilisation the seed coat was nicked and left to imbibein sterile water for at least 2 h. To facilitate even germi-nation, the seed coat and endosperm were ascepticallyremoved and the germinating seedling placed on 0.8%agar. After 1 week the shoot tips of the seedlingswere transferred to CR medium without hormones[31] whilst the roots were harvested for the histochem-ical GUS assay. Plantlets with roots formed after 3to 4 weeks of culture. After an additional 4 weeks,plantlets were transferred to sterile soil and establishedin a containment glasshouse.

β-glucuronidase assays

GUS activity was determined using both histochem-ical and fluorometric assays. For the histochemicalassay tissue segments were incubated overnight at37 ◦C in 1 mg/ml 5-bromo-4-chloro-3-indolyl-β-D-glucuronide (X-GlcA CHX) in 50 mM sodium phos-

phate buffer pH 7.0, 0.1% (v/v) Triton X-100 and 20%(v/v) methanol. Histochemical assays were performedon the root and part of the hypocotyl of 1 weekin vitrogrown seedlings, roots and leaf segments of 3-4 weekin vitro grown plantlets and root tips, internode stolonsections and leaf segments of glasshouse-establishedplants. A leaf segment consisted of the portion of thelamina adjacent to the petiole, taken from the youngestfully expanded leaf. Leaf segments were harvestedfrom at least 5 different stolons. The fluorometric GUSassay was performed as described by Jefferson [17] us-ing 4-methylumbelliferyl-β-D-glucuronide (MUG) asthe substrate. Assays were conducted on mature plantsestablished in the glasshouse. For each plant assayed,leaf sections were pooled from the terminal tip of theyoungest fully expanded leaf from 5 different stolons.

PCR

A modified procedure of Edwardset al. [11] was usedto extract DNA for PCR analysis. Leaf tissue equiva-lent in size to an Eppendorf tube lid was frozen withliquid nitrogen then ground with a disposable grinder.A 400µl portion of extraction buffer (200 mM Tris-

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HCl pH 7.5, 250 mM NaCl, 25 mM EDTA, 0.5%SDS) and 400µl phenol were added to the macer-ated plant material and the sample vortexed for 5 s.Once all the samples had been collected, the extractswere centrifuged at 13000 rpm for 5 min. 300µlof supernatant was transferred to a fresh Eppendorfand mixed by inversion with an equal volume of iso-propanol, then left at room temperature for 2 min.After centrifugation at 13000 rpm for 5 min, the pel-let was vacuum-dried and resuspended in 50µl ofsterile water. DNA amplification occurred in 50µlof reaction mix containing 160µM of each dNTPs,800 nM of eachuidAprimer (5′-CAGGAAGTGATG-GAGCATCAGG-3′, 5′-CAGTCGAGCATCTCTTCAG-CGT-3′), 5 µl PCR 10 × Reaction Buffer+Mg(Boehringer Mannheim), 5µl of extracted DNAand 2.5 units ofTaq DNA polymerase (BoehringerMannheim). Ampliwax (Perkin Elmer) was used toseparate dNTPs and primers from the buffer and DNA.The amplification was performed in a Perkin Elmer480 thermocycler. The reaction mixture was held at92 ◦C for 2 min then proceeded for 40 cycles underthe following conditions: 92◦C for 60 s, 55◦C for60 s and 72◦C for 100 s, terminating with a 72◦C10 min extension for the last cycle.

Southern hybridisation

DNA from the PCR gel was capillary blotted onto anylon membrane (Hybond N+, Amersham) and hy-bridised according to manufacturers’ instructions with[32P]-labelled 1900 bpEcoRI/NcoI fragment frompJIT166 [15], prepared using a random primer proce-dure (Ready-to-Go, Pharmacia). The probe containeda promoterlessuidAgene.

EcoRI, NcoI, EcoRI/NcoI genomic DNA digests ofprimary transgenic and progeny plants were made andSouthern- blotted according to Voiseyet al. [28]. Thedigested DNA was probed withuidA gene fragmentfrom pJIT166 and with a 536 bpNcoI/SphI frag-ment from pANDY1 (Andrew Griffiths, unpublished)containing the promoterlessnptII gene.

Results

BC1 and BC2 inheritance

Results of the segregation analysis for the BC1 andBC2 populations, using the GUS histochemical as-say are presented in Table 1. No significant differencein GUS segregation was observed between reciprocal

Figure 2. PCR amplification of a 715 bp ofuidA fragment fromGUS-negative (lanes 1-8), positive F2 plants (lane 9) and a controlplasmid (lane 10) containing theuidA gene.

crosses in the BC1 population. Since a maternal ef-fect was not detected, subsequent crosses were madeonly in one direction. The segregation ratios obtainedfor the BC1 population and both the low and highGUS expression subsets of the BC2 population did notsignificantly deviate from that expected for a singledominant gene.

F2 inheritance

Individual plants within the F2 population were his-tochemically assayed for GUS expression (Table 2).Segregation ratios for individual crosses within the F2population varied. For both Cross 1 and 3, the GUSsegregation ratio did not differ from that expected fora single dominant locus. Whereas in Cross 2, the seg-regation ratio obtained (1.2:1) did not fit the expected3:1 ratio (χ2

1df P< 0.01). For each cross, reserve seedwas germinated and assayed for GUS activity (seeSeed Batch 2, Table 2). The segregation ratios ob-tained were similar to that observed for the first batchof seed. Again the same aberrant GUS segregation ra-tio was observed for Cross 2. For both batches of seed,the segregation ratios obtained did not differ from thatexpected (2:1) for a recessive lethal (χ2

1df P > 0.05).The distortion in the F2 segregation ratio observed

in Cross 2 was not caused by inactivation of theuidAtransgene. All the GUS-negative plantlets in the sec-ond batch of seed were screened for the presence of theuidA transgene using PCR techniques (see Figure 2).Amplification of the 715 bpuidA gene fragmentdid not occur in any of the GUS-negative F2 plantsduring PCR experiments. Therefore, none of the GUS-negative F2 plants had inherited an inactivateduidAtransgene. Southern hybridisation experiments on di-gested genomic DNA indicated that no detectable re-

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Table 1. Segregation of theuidA transgene in BC1 and BC2 populations. Histochemical staining was usedto determine GUS activity in 1-week old seedlings. NS = not significant (P > 0.05).

Population Cross GUS-positive GUS-negative Deviation from 1:1 ratioχ21df

BC1 GH7× GH9 17 19

GH9× GH7 11 19

total 28 38 NS

BC2H GH646× C6525/13 13 14

GH652× C6525/13 5 3

GH694× C6525/13 8 3

GH708× C6525/13 7 5

total 33 25 NS

BC2L GH637× C6525/13 3 3

GH698× C6525/13 9 5

GH644× C6525/13 11 11

GH633× C6525/13 6 4

total 29 23 NS

Table 2. Segregation of GUS activity in F2 populations based upon histochemical staining of 1-weekold seedlings.

Seed Batch 1 Seed Batch 2

Cross Positive Negative Positive Negative

1 (GH747× GH646 and GH646× GH747) 20 (21) 8 (7) NS 35 (33) 9 (11) NS

2 (GH640× GH642) 18 (25) 15 (8)∗∗ 20 (27) 16 (9)∗∗3 (GH694× GH646) 10 (11) 5 (4) NS 6 (10.5) 8 (3.5)∗∗

Total 48 28∗ 61 33∗Figures in parenthesis are expected cell values based upon a 3:1 positive/negative ratio. Deviation fromexpected 3:1 ratio: NS, not significant;∗ P < 0.05;∗∗ P < 0.01,χ2 test l df.

arrangement of the T-DNA had occurred between theprimary transgenic and progeny plants (Figure 3). Amuch largerHindIII fragment than the expected 4.2 kbband was obtained in the primary transgenic plant andsubsequent progeny (Figure 3) although often a faint4.2 kb band could be seen. It is possible that theHindIII site at the end of theuidA ocs 3′ was presentbut was failing to cut to completion. The adjacentEcoRI site and the otherHindIII within the T-DNAboth cut to completion. The banding patterns obtaineddid not differ from plasmid digests of the constructoriginally used to transform the parent plant. Thusprogeny and the original primary transgenic plant hadan intact copy of the T-DNA. Therefore the segrega-tion distortion seen in Cross 2 was not due to a partialdeletion of theuidA transgene.

Separation of homozygous from heterozygous classes

Since F2 plants that are either heterozygous or ho-mozygous for theuidA transgene both exhibit GUSactivity, it was necessary to test cross individual plantsto a non-transgenic control to identify the two groups.In the heterozygous group, the transgene occurs inonly one of the two chromosomes in a homologouspair. Therefore theuidA gene is transmitted to onlyhalf of the progeny whereas in a homozygous plantall the progeny would be expected to inherit the trans-genic trait. A total of 37 GUS-positive F2 plants wereprogeny-tested and the GUS segregation ratios ob-tained for these families are presented in Table 3. Foreach Cross, the number of F2 plants homozygous orheterozygous for theuidA transgene is summarised inTable 4. In Crosses 1 and 3, the segregation ratiosobtained (homozygous/heterozygous) did not signifi-

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Table 3. GUS segregation ratios obtained after test crossing GUS-positive F2 plants.

F2 plant Non-transgenic tester Segregation ratio Deviation from F2 plant status

positive/negative 1:1 ratio

Cross 1b1 × I2/18 11 9 NS Heterozygous

b7 × I2/18 20 0 ∗ ∗ ∗ Homozygous

b9 × I2/6 10 10 NS Heterozygote

c2 × I2/6 11 9 NS Heterozygous

c3 × I2/6 20 0 ∗ ∗ ∗ Homozygous




c10 × I2/18 20 0 ∗ ∗ ∗ Homozygous

d2 × I2/6 8 12 NS Heterozygous





d8 × I2/18 4 16 ∗ Heterozygous

Cross 2e8 × I2/18 9 11 NS Heterozygous

f1 × I2/18 12 8 NS Heterozygous









g1 × I2/6 12 8 NS Heterozygous


g3 × I2/6 20 0 ∗ ∗ ∗ Homozygous


Cross 3h1 × I2/18 8 12 NS Heterozygous

h4 × I2/18 20 0 ∗ ∗ ∗ Homozygous

h7 × I2/18 9 11 NS Heterozygous

i1 × I2/6 10 10 NS Heterozygous

i2 × I2/18 20 0 ∗ ∗ ∗ Homozygous

i3 × I2/18 19 1 ∗ ∗ ∗ Homozygous

i5 × I2/6 13 7 NS Heterozygote

i7 × I2/18 5 15 ∗ Heterozygous

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Figure 3. Southern analysis of primary transgenic and progenyplants. DNA was isolated and hybridised with the GUS coding re-gion. A. Restriction Map of the T-DNA region of plasmid pPE64used in transformation. B. Genomic DNA from each plant (inthis instance GH647) was digested with a range of restriction en-zymes (lane 1,HindIII; lane 2, NcoI/EcoRI; lane 3,BglII; lane 4,BglII/HindIII; lane 5,EcoRI; lane 6,NcoI; lane 7,Bg1II/HindIII. C.The restriction patterns obtained, in this instance withNcoI diges-tion, indicated that no rearrangement of the T-DNA had occurred be-tween the primary transgenic plant and progeny plants. Lanes 1–6contain GUS-positive progeny (GH646, GH642, GH1388, GH1452,GH694 and GH647), lane 7 the non-transgenic parent (GH9) andlane 8 the primary transgenic plant (GH7).

cantly differ from that of the expected 1:2. In Cross 2there was a significant decrease in the expected num-ber of homozygous plants obtained (ratio of 1:13). Thedistortion in this ratio did not result from misclassifi-cation of homozygous as heterozygous plants since atleast three GUS-negative progeny from each heterozy-gous plant were analysed by PCR for the presence ofthe uidA transgene. No cases of transgene inactiva-tion were detected among the GUS-negative progenyscreened.

Test cross progeny analysis

Due to a delay in the onset of GUS activity, seg-regation results were based upon the second GUShistochemical assay. The majority of progeny seriesfrom the F2 plants segregated as expected. Either all20 progeny had GUS activity or the ratio of GUS-positive to GUS-negative plants was not significantlydifferent from the expected 1:1 (χ2

1df P > 0.05). How-ever, unexpected segregation ratios were obtained inthe progeny from some F2 plants (see Table 3).

In Cross 3, progeny from F2 plant i3 segregated19:1 (positive/negative) for GUS activity. PCR analy-sis withuidA andnptII primers and Southern hybridi-sation of genomic DNA revealed that the single GUS-negative plant lacked both theuidA andnptII genes.The segregation ratio significantly differed from thatexpected of a heterozygous plant (χ2

1df P < 0.001).A further 20 progeny from plant i3 were assayed forGUS activity. All were GUS positive. Therefore planti3 was regarded as being homozygous for the T-DNAinsert.

Two other plants, d8 and i7, from Crosses 1 and 3,also had GUS segregation ratios that did not fit the ex-pected 1:1 ratio. In both these plants there were morethan the expected number of GUS-negative progeny.In these instances the distortion in the segregation ratiowas not due to inactivation of the transgene since noneof the GUS-negative progeny from these plants had in-herited theuidAgene as judged by PCR. An additional20 seeds from plant d8 were assayed for GUS activ-ity and did not significantly differ from the expected1:1 ratio (12 negative/ 8 positive). This result is notsurprising given that the number of cases with unex-pected segregation ratios (2/37) were equivalent to theexpected type 1 error rate of 5%. Progeny from themajority of heterozygous plants did not appear to beaffected by transgene inactivation. This is reflected inthat the overall segregation ratio obtained for progenyfrom heterozygous F2 plants (265 GUS-positive and

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Table 4. Segregation of homozygous and heterozygous plants in F2 populations.

Cross Homozygous plants Heterozygous plants Totalχ2l df (1:2)

1 3 12 15 NS

2 1 13 14 P < 0.05

3 3 5 8 NS

Total 7 30 37

Expected value 12.33 24.67 37 0.10> P > 0.05

1:2 ratio

295 GUS-negative), although containing a slight ex-cess of negative plants, was not significantly differentfrom the expected 1:1 ratio (χ2

1df P > 0.05).

Delayed onset of GUS activity

In the F2 population the results of subsequent GUShistochemical assays on individual plants were sta-ble across developmental age. Consistent assay resultswere obtained whether the plant was a one week oldseedling, a plantlet grown in tissue culture for fourweeks or an established glasshouse plant. The majorityof plants in the BC1 and BC2 populations also ex-hibited stable developmental GUS activity. However,three plants subsequently developed GUS activity be-tween the first histochemical assay on one-week oldseedlings and the second assay on plantlets grown forthree to four weeks in tissue culture.

This delay in the onset of GUS activity was verymarked in the test cross progeny, where 22% (n=340)of the plants tested subsequently developed GUS ac-tivity between the first and second histochemical test;42% of plants exhibited GUS activity on the firsthistochemical assay. Often the intensity of GUS his-tochemical staining increased between the first andsecond tests.

The delay in onset of GUS activity was seen inCrosses 1, 2 and 3 and in progeny derived from bothheterozygous and homozygous plants. Within eachcross, there were F2 plants where none of the progenywere affected by the delay in onset of GUS activity.

Quantification of GUS activity

GUS activity was quantified in the BC1, BC2 andF2 populations. The median values obtained for theBC1 and BC2 populations were lower than that of theoriginal primary transgenic. This reflected the consid-erable variation in GUS activity found within these

Figure 4. Quantification of GUS activity. Data are represented asbox plots [7]. Horizontal bars indicate the 10th, 25th, 75th and90th percentiles. Outliers are marked by an open circle or an aster-isk. BC1, primary transgenic× non-transgenic plant; BC2, BC1 ×non-transgenic plant; F2, BC1 × BC1; N, GUS activity of primarytransgenic plant (GH7).

populations. Though individuals were identified withthe same mean GUS activity as that of the primarytransgenic, others were found with significantly lowerGUS activity (Figure 4). No difference was found inGUS activity amongst the BC1, BC2 and F2 popula-tions derived from GH7 for GUS-positive individuals.GUS expression was also quantified in populations(BC2H ) derived from the three highest expressingBC1 plants and populations (BC2L) derived from thethree lowest-expressing BC1 plants (see Figure 5). Theoverall mean of the BC2H population (5356± 1335)was significantly higher than that of the BC2L pop-ulation (3633± 928). No significant difference wasdetected among the BC2H populations or among theBC2L populations. GUS expression for known het-erozygous and homozygous individuals was quantifiedusing a fluorometric assay. The mean GUS activity

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Figure 5. Quantification of GUS activity between BC2H and BC2Lpopulations. Populations are represented as box plots. BC2H pop-ulations (numbers 1–3) were derived from plants with high GUSactivity (GH646, GH652, GH708) whilst BC2L populations (num-bers 4–6) were derived from plants with low GUS activity (GH698,GH644, GH633).N, GUS activity of GUS-positive parent plant.

Figure 6. Box plot comparison of GUS activity in heterozygous(n=30) vs. homozygous plants (n=7) within the F2 population.

for the homozygous population though slightly higherthan that of the heterozygous population, was notsignificantly different (Figure 6).

Discussion

We investigated the effects of allelic composition andgenetic background on the expression and inheritanceof a CaMV 35Sp:uidA transgene in white clover. Wehave demonstrated that a singleuidA insert can be sta-bly expressed and transmitted in a Mendelian mannerto white clover progeny in either a heterozygous or ho-

mozygous state. However, non-Mendelian inheritancewas also observed in one F2 population.

There was no significant difference in the me-dian values obtained for GUS expression among theBC1, BC2 and F2 populations. This indicated that theuidA transgene was stably expressed from one gen-eration to another. Stable expression and inheritanceof a transgene has also been observed in alfalfa [21]and in maize [13] across successive generations. Con-siderable (four fold) variation in the level of GUSexpression was observed among plants within eachpopulation, which is similar to the level of varia-tion observed by Fearinget al. [13], who obtaineda 10 fold difference among progeny expressing aCryIA(b) transgene. Variation in expression levelsamong doubled haploid lines derived from the samehaploid transgenic plant also has been observed [25].This variation in GUS activity was probably relatedto the heterogeneous genetic makeup of each individ-ual plant. White clover is an outbreeding species sothat considerable genetic variation exists between evenclosely related individuals. Thus although each plantin the backcross population contained one copy of thesame T-DNA insert, the T-DNA insert was effectivelyin a different genetic background. The differences ingenetic background may be affecting the expressionof the uidA transgene thus contributing to the con-siderable variation in GUS activity observed withinpopulations. In this regard, the variation seen betweenindividuals within the same population was similarto the variation observed among primary transgenicplants transformed with the same construct.

Crossing experiments involving high and lowGUS-expressing plants indicated that GUS expres-sion was influenced by the genetic background of theplant and thus it is possible to select for transgeneexpression. This variation emphasises the need in anoutbreeding species to test transgenic expression andstability in a variety of genetic backgrounds beforeproceeding to a breeding programme.

The number of GUS negative plants was over-represented in progeny from Cross 2. An excess of thenon-transgenic class in inheritance studies has beenwidely reported in the literature. In the majority ofinstances where this phenomenon has been investi-gated at the molecular level, the under-representationof the transgenic class was found to be due to trans-gene inactivation [14]. This was not the case in thisstudy, since PCR and Southern analysis demonstratedthat none of the GUS-negative plants from this crosshad inherited an inactivateduidA transgene. It is in-

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teresting to speculate why the same transformationevent should be inherited in a Mendelian manner inone genetic background and not in another. Meioticinstability of transgenes resulting in non-Mendelianinheritance has been observed in some studies [5, 6,27]. No difference in the Southern hybridisation band-ing pattern for theuidA andnptII genes was detectedamong the primary transgenic, BC1 and F2 plants.This suggests that the segregation distortion seen inthe F2 population was not due to re-arrangement ofthe T-DNA. The segregation ratio of GUS-positive toGUS-negative plants in Cross 2 did not fit the expected3:1 ratio for a single dominant gene under Mendelianinheritance but fitted the 2:1 ratio expected for a reces-sive lethal. The presence of a plant homozygous forthe reporter gene in this cross, indicated that the trans-gene itself was not functioning as a recessive lethal,although the number of plants homozygous for thetransgene was less than expected. The reduction inthe homozygous class alone was insufficient to fullyaccount for the distortion seen in the expected 3:1 ra-tio of GUS-positive to GUS-negative plants. Givena 1:13 segregation of homozygous to heterozygousplants, the ratio obtained foruidAhomozygous to het-erozygous to non-transgenic plants was 1:11:10. Thus,though not to the same extent, the heterozygous classwas also under-represented in this cross.

Peng et al. [24] demonstrated stable transmis-sion of a transgene from one generation to another intransgenic lines of rice but reported distorted segre-gation ratios in two out of three lines in F2 crossesand were unable to obtain plants homozygous for thetransgene from these lines. Others have also obtaineddistorted transgene segregation ratios that could notbe explained by transgene inactivation [26, 29]. Bothgroups were working with inbred lines of maize. Theauthors suggested several reasons to explain the trans-gene segregation distortion observed; including theincorporation of the transgene into an essential gene,that the site of gene insertion could affect gameteviability, or that the introduced DNA is unstably in-tegrated resulting in the absence of the transgene insome of the gametes.

An alternative hypothesis is that the segregationdistortion is not due to the immediate site of insertionor the transformation event, but is a consequence ofthe genetic background. White clover is an outbreed-ing species, with a high degree of heterozygosity andthus can tolerate a high genetic load of deleteriousrecessive lethals [34]. It is possible that the trans-gene has inserted near a recessive lethal. Generally

plants homozygous for the transgene also would behomozygous for the recessive lethal and thus non-viable. Crossing over events between the transgeneand the recessive lethal would account for the recoveryof the occasional plant homozygous for the transgeneand the subsequent under-representation of plants inthe heterozygous class.

Instances of non-Mendelian segregation involvingnon transformed plants have been reported in a widerange of plant species [18]. Bradshaw and Stettler [3]observed segregation distortion of a marker locus re-sulting from the tight linkage of the locus to a recessivelethal allele. Thus aberrant segregation ratios due togenetic load are already a factor in the breeding ofoutcrossing species. In this respect handling distortedtransgene segregation ratios in cultivar developmentis no different to problems already encountered intraditional breeding.

The occurrence of segregation distortion in a spe-cific F2 population but not in backcross populations,emphasises the need in outbreeding species to testtransgenes in a range of genetic backgrounds and in ahomozygous state. This view is reinforced by Neuhu-beret al. [23] who found that the instability of specifictransgene loci was not initially apparent in the origi-nal primary transgenic, nor in backcross populations,but was only revealed when the tobacco plants werepropagated for several generations as homozygotes.

Usually all the progeny of a homozygous trans-genic plant would inherit the transgenic trait. Plant i3differed from expectation in that a single progeny plantthat lacked theuidA and nptII genes was obtained.This plant may have resulted from a rare meiotic in-stability event that led to the loss of part or all ofthe T-DNA, though the possibility of accidental seedcontamination cannot be entirely eliminated. Low fre-quencies of meiotic transgene instability have beenreported for plants transformed viaAgrobacteriumwith single-copy inserts [6, 22] whilst other work-ers [5] have documented the actual loss of transgenesequences from one generation of plants to another.

The delay in the onset of gus activity seen in thisstudy was similar to the expression pattern obtainedfor domain A (−90 to+8) of the 35S promoter byBenfeyet al. [1]. These authors found in transgenictobacco plants [1, 2] that developmental regulationof expression patterns was influenced by a series ofsubdomains within the 35S promoter. Root expressionwas mainly determined by Domain A. Expression wasobserved in the roots of ten day old seedlings but notof six day old plants, for individuals with only Domain

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A. When both domains A and B were present, domainB (−343 to−90) modified the expression pattern seenso that root expression was detected earlier. The 35Spromoter used to drive theuidAgene in this study wastruncated (−388 to+2) and lacked the B5 subdomain.

In white clover, the delay in onset of gus activitydiffered between different populations, even thoughall populations were derived from the same initialtransformation event. Thus the genetic backgroundof the plants may be influencing the developmentalregulation of the 35S promoter.

In this study the heterozygous plants contain twocopies of the CaMV 35S promoter, one transcribingthe uidA transgene and the other controlling the se-lectable marker. Therefore, homozygous transgenic F2plants contain four copies of the CaMV 35S promoter.This can be disadvantageous since it has been foundthat multiple copies of a gene are more prone to ge-netic inactivation than single copies [16]. Despite thepresence of four copies of the CaMV 35S promoter,we found no evidence ofuidAgene inactivation amongF2 plants homozygous for this transgene.

We obtained homozygous GUS-positive plants thatexpress at a similar level to that of heterozygousplants. Caligariet al. [4], Penget al. [24] and Fearinget al. [13] also found no difference in GUS activ-ity between transgenic homozygous and heterozygousplants, whereas others [8, 16, 20] have found increasedactivity of a single-copy transgene in a homozygousstate. Conversely, Elmayan and Vaucheret [12], usinga strongly expressed 35SuidA gene, found that GUSexpression was silenced in homozygous plants.

The ability to select high-expressing transgenicplants homozygous for an introduced gene is a consid-erable advantage in breeding programmes. In a planthomozygous for the transgene, the introduced geneis fixed so that all progeny from this plant inherit thetransgene [34]. It is therefore possible to intercross aprimary transgenic plant to the parent plants or a rangeof plants of an existing white clover cultivar and se-lect for high-expressing homozygous transgenic plantsfrom each line. These individuals can then be inter-crossed so that the transgenic trait can be fixed withinthe cultivar. The new cultivar would still retain thehigh degree of genetic diversity present in the originalcultivar. Here we have demonstrated the stable expres-sion and inheritance of a transgene in an outbreedingspecies as well as the importance of determining theeffects of genetic background on transgene expressionand transmission.

Acknowledgements

We thank Dorothy Maher and Anne Allan for theirexpert technical assistance and Dr Ian Henderson forstatistical advice. We are grateful for the support andencouragement of all members of the Plant Molecu-lar Genetics Laboratory. This work was funded by theNew Zealand Foundation for Research, Science andTechnology, Grant C10 405 (D.W.R.W.).

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