Theoretical and Applied Genetics

, Volume 118, Issue 2, pp 259–273

Association mapping reveals gene action and interactions in the determination of flowering time in barley

  • Silke Stracke
  • Grit Haseneyer
  • Jean-Baptiste Veyrieras
  • Hartwig H. Geiger
  • Sascha Sauer
  • Andreas Graner
  • Hans-Peter Piepho
Open AccessOriginal Paper

DOI: 10.1007/s00122-008-0896-y

Cite this article as:
Stracke, S., Haseneyer, G., Veyrieras, JB. et al. Theor Appl Genet (2009) 118: 259. doi:10.1007/s00122-008-0896-y

Abstract

The interaction between members of a gene network has an important impact on the variation of quantitative traits, and can influence the outcome of phenotype/genotype association studies. Three genes (Ppd-H1, HvCO1, HvFT1) known to play an essential role in the regulation of flowering time under long days in barley were subjected to an analysis of nucleotide diversity in a collection of 220 spring barley accessions. The coding region of Ppd-H1 was highly diverse, while both HvCO1 and HvFT1 showed a rather limited level of diversity. Within all three genes, the extent of linkage disequilibrium was variable, but on average only moderate. Ppd-H1 is strongly associated with flowering time across four environments, showing a difference of five to ten days between the most extreme haplotypes. The association between flowering time and the variation at HvFT1 and HvCO1 was strongly dependent on the haplotype present at Ppd-H1. The interaction between HvCO1 and Ppd-H1 was statistically significant, but this association disappeared when the analysis was corrected for the geographical origin of the accessions. No association existed between flowering time and allelic variation at HvFT1. In contrast to Ppd-H1, functional variation at both HvCO1 and HvFT1 is limited in cultivated barley.

Introduction

The timing of flowering is a key component of plant adaptation and so represents a major objective for crop breeding. Although some major genes have been identified in the control of flowering time in crop species (Cockram et al. 2007; Hay and Ellis 1998; Laurie 1997), the molecular dissection of flowering time in the leading plant model Arabidopsis thaliana has shown that a complex network of regulatory pathways is involved (reviews: Ausin et al. 2005; Hay and Ellis 1998; Henderson and Dean 2004; Imaizumi and Kay 2006; Mouradov et al. 2002; Putterill et al. 2004). Among the crop species, the best understood system is rice, where five heading date genes (Hd1Hd5) have been successfully tagged or isolated, and the DNA structures of these genes show a remarkable level of similarity to those in A. thaliana (Kojima et al. 2002; Lin et al. 2003; Yamamoto et al. 1998; Yano et al. 2000, 2001). Two environmental cues are involved in the determination of the flowering time of barley. One of these, temperature, acts through the vernalisation pathway; the other is daylength, acting through the photoperiod sensitivity Ppd genes. One of the latter, Ppd-H1, has recently been cloned (Turner et al. 2005). As it represents a homolog of A. thalianaPRR7, it may be involved in the functioning of the circadian clock (Nakamichi et al. 2005). The barley homologs of CONSTANS (HvCO1, 7HS; Griffiths et al. 2003) and FLOWERING LOCUS T (HvFT1 synonym of Vrn-H3, 7HS; Faure et al. 2007; Turner et al. 2005; Yan et al. 2006) both act downstream of Ppd-H1, 2HS (Turner et al. 2005). In rice, these genes are represented by Hd1 and Hd3a, respectively. In A. thaliana, the circadian clock regulated CO encodes a B-box-type zinc finger transcriptional activator (Putterill et al. 1995), which interacts with a transcription factor complex to induce the expression of FT (Ben-Naim et al. 2006; Cheng and Wang 2005; Kardailsky et al. 1999; Suarez-Lopez et al. 2001). While the CO gene product acts exclusively in the photoperiodic pathway (Yanovsky and Kay 2002), the FT product is involved in other flowering pathways as well (Abe et al. 2005; Ausin et al. 2005; Kardailsky et al. 1999; Wigge et al. 2005).

The identification of the three genes Ppd-H1, HvCO1, and HvFT1 as players in the determination of flowering time in barley makes them attractive as candidates for a phenotype/genotype association analysis across a wide germplasm sample. In this study we set out to search for associations between nucleotide diversity at these three genes and variation in flowering time within a geographically widely distributed collection of spring barleys. The multi-factorial inheritance of most quantitative traits complicates their genetic analysis. Although the generally adopted assumption is that quantitative trait loci (QTL) act additively, significant QTL × QTL interactions have been observed across a range of plant species (Alonso-Blanco et al. 1998; Caicedo et al. 2004; Eshed and Zamir 1996; Juenger et al. 2005; Li et al. 1997, 2000; Lukens and Doebley 1999). We have analyzed the present data set to examine the impact of statistical gene × gene interactions on the outcome of association results.

Materials and methods

Plant material

The germplasm collection consisted of 220 spring barley accessions originating from Europe (EU, n = 108), East Asia (EA, n = 39), West Asia and North Africa (WA, n = 44), and America (AM, n = 29) (Supplemental Table S1). The accessions are a subset of the full Barley Core Collection (“BCC”; Knüpffer and Hintum 2003), chosen to represent the widest possible range of diversity, with some additional West Asian accessions (“HOR”) provided by the Gatersleben Genebank.

Phenotypic evaluation

Spring sown field trials were conducted at three locations, two in Southern (Stuttgart-Hohenheim and Irlbach) and one in Northern (Bergen-Wohlde) Germany. Flowering time was recorded in 2004 at Irlbach and at all three sites in 2005. Each trial was arranged in microplots in a 15 × 25 lattice design, with three replicates. Flowering date was recorded as days after sowing (das), using the date at which half of the ear had emerged from the flag leaf of the main tiller in 50% of the plants within a given plot.

Genotypic analysis

Genomic DNA was extracted from pooled leaf tissue of six plants per accession, as described elsewhere (Graner et al. 1991). Twenty-five barley expressed sequence tag-derived simple sequence repeat markers (EST-SSRs) (listed in Supplemental Table S2; Thiel et al. 2003) were used to characterize population structure. The criteria applied for marker selection were map position (to maximise genome coverage) and informativeness. The SSRs were grouped into multiplex sets of three to six each, and amplicons were generated with the Multiplex PCR kit (QIAGEN), using an amplification program of 95°C/15 min, followed by 40 cycles of 94°C/30 s, 60°C/30 s and 72°C/15 s, and a final extension step of 72°C/10 min. PCR products were separated on 6% polyacrylamide gels on an ABI377 system (Applied Biosystems/Applera, Darmstadt, Germany), and profiles were analyzed with GeneScan 3.7.1 und Genotyper 3.7 software.

Sequence analysis

Primers were designed to amplify coding and non-coding sequence intervals of the candidate genes, covering 1230 bp (Ppd-H1), 1772 bp (HvCO1) and 1974 bp (HvFT1). The genomic position of each amplicon was inferred from the respective reference sequence (Ppd-H1–AY970701, HvCO1–AF490467, and HvFT1–DQ100327). Primer sequences and relevant reaction conditions are given in supplemental Table S3. The Ppd-H1 fragment includes the region from intron 4 to the beginning of exon 7, and covered the entire exon 5 and 6 as well as the entire intron 5 and 6. The region contains one of the four base pair positions diagnostic for the early and the late flowering alleles Ppd-H1 and ppd-H1 (Turner et al. 2005). The HvCO1 sequence covered most of the gene, with small gaps in the first intron and at the 5′ and 3′ ends. HvFT1 was resequenced in its entirety, except for 100 bp at its 3′ end. As FT regulates flowering time in a dosage-dependent manner (Kardailsky et al. 1999), an additional 1 kbp of the 5′ upstream region was included. In preparation for sequencing, DNA fragments were amplified using AccuPrime DNA polymerase (Invitrogen). The PCR amplicons were purified in 384 well plates and their concentrations were equalized. Ten nanogram PCR product served as the template for cycle-sequencing using ABI BigDye Terminator 3.1 chemistry on a capillary automatic sequencing device (3730xl ABI 96; Applied Biosystems/Applera, Darmstadt, Germany). The amplification primers were used as sequencing primers. For quality scoring a software package based on the poly-phred system was used (Nickerson et al. 1997). Alignments were compiled and analyzed using Sequencher v4.1 (Gene Codes Corporation, Ann Arbor, USA) and Bioedit v4.7.8 (http://www.mbio.ncsu.edu/BioEdit/bioedit.html).

Population structure

Population structure was analysed in two ways. First, within a Bayesian framework, a Markov chain Monte Carlo algorithm was used to sample from the joint posterior distribution of the subpopulation allele frequencies, and assignment of individuals to particular sub-populations was effected with STRUCTURE version 2.1 (Pritchard et al. 2000a; http://pritch.bsd.uchicago.edu/structure.html). QST is the matrix of admixture proportions obtained with STRUCTURE. For between two and ten subpopulations, ten independent simulations were performed, using the admixture model and a burn-in of 500,000 followed by 5,000,000 iterations. The ‘optimal’ number of subpopulations K was chosen on the basis of the second order rate of change of the likelihood function with respect to K (Evanno et al. 2005). The second approach has been developed by Veyrieras et al. (2006) from the Frequentist framework devised by Tang et al. (2005). For a given value of K, an expectation maximization (EM) algorithm was run with ten random starting points to obtain the maximum likelihood estimates of both the admixture proportions and the population allele frequencies. Only the run with the highest likelihood was retained. For each value of K, the cells of the allele count matrix C (Cij = 1 if individual i carries allele j, 0 otherwise) were adjusted according to their expected frequencies under the admixture model with K populations (i.e., the estimated individual admixture proportion was multiplied by the allele frequencies in each population). Then the spectral norm of the resulting adjusted matrix M (i.e., the eigenvalue of the covariance matrix MM) was used as a measure of the linkage disequilibrium (LD) between marker alleles not explained by the current admixture model. Spectral norms are expected to decrease until reaching a plate after the ‘optimal’ value of K. The model selection is thus based on a scree plot of these spectral norms, in order to identify the value of K after which the decrease is no longer significant (see Supplemental Figure S1). This value is referred to as KEM, and the associated matrix of admixture proportions as QEM.

Allele diversity

All cultivars were treated as pure lines and the small proportion of non-homozygous data points was treated as missing data. For the purpose of statistical analysis, indels were regarded as single site polymorphisms. Only haplotypes represented by at least two accessions were considered. To assess nucleotide variability across the candidate sequences, and to measure and compare the diversity within and across geographical origins, both nucleotide (pi) and haplotype (hd) diversities were estimated using DnaSP (Version 3.51; Rozas and Rozas 1999; http://www.ub.es/dnasp/). Estimates for LD were obtained with TASSEL software (v1.0.9; http://www.maizegenetics.net/bioinformatics/tasselindex.htm) applying the measurement r2 (squared correlation coefficient; Hill and Robertso 1968) and the significance of any LD was determined using Fisher’s two-tailed exact test. Values of r2 were plotted against the pairwise distance between polymorphic sites, or presented in a matrix plot.

Association analysis

For the analysis of phenotypic data, the adjusted means per environment were first computed, and these were subjected to a mixed model analysis over trials. Individual trials were analyzed by a mixed model using fixed effects for genotype and replicate, and random block effects for recovery of information. Variance components were estimated by the REML method. For each trial, adjusted means and weights were computed from the diagonal elements of the inverse of the asymptotic variance–covariance matrix of adjusted means (Smith et al. 2001). In the subsequent step, the following terms were fitted: trial main effect (fixed), genotype main effect (random), genotype-by-trial interaction (random). While our final model assumes random genotypes, we initially take the genotypic effect as fixed in the first step. This is necessary because taking genotypes as random already in the first step we would be estimating genotype means by best linear unbiased prediction (BLUP), which involves shrinkage. The degree of shrinkage usually varies among trials, and so BLUPs would be differently scaled and therefore not comparable across trials. For this reason, BLUPs cannot be used as data in stage two (Smith et al. 2001). Taking genotypes as fixed in stage one, we obtain unbiased estimates of genotype means, for which the model in stage two is (approximately) valid. In addition, the population structure was modeled by fixed-effects regression on an N × K matrix Q of probabilities of membership for K sub-population membership and each of N genotypes. The Q matrix was either computed using the approach of Pritchard et al. (2000a) or Veyrieras et al. (2006) as described above. For model comparison, two approaches were followed to compute a pairwise kinship matrix using the 25 background SSRs. One approach was equivalent to ridge regression for the allele effects. We generated a dummy variable for each allele taking value unity when the allele was present and zero when absent. For each dummy a random linear regression coefficient was fitted. Coefficients for all alleles were assumed to be sampled from a common normal distribution. The model was fitted by using a Toepitz structure as implemented in the MIXED procedure of the SAS system. It corresponds to fitting a linear variance structure with a fixed matrix corresponding to a kinship matrix. For further details, including implementation in the MIXED procedure see Ruppert et al. 2003). The second kinship matrix was computed based on estimated Nason’s coefficients (Loiselle et al. 1995) using SPAGeDI version 1.2 software (Hardy and Vekemans 2002). Negative kinship values were set to zero following Yu et al. 2006). Each kinship matrix was fitted for random genotype main effects and random genotype-by-trial interaction effects. Model selection with regard to admixture probabilities (Q) and kinship was performed using Akaike’s Information Criterion (AIC) in “smaller-is-better” form. All these analyses used components of PROC MIXED in the SAS System (v9.1.3) package. ML was used instead of REML for model selection to enable comparison between models with different fixed effects structures. Tests of fixed effects based on selected models were then based on the REML method with denominator degrees of freedom approximated by the Kenward-Roger method. Adjusted means were compared by Wald t tests. The family-wise type I error rate was controlled by the Bonferroni–Holm procedure. As the means were not variance balanced, we used the method of (Piepho 2004) to generate a letter display showing the significance of comparisons. In order to identify gene × gene interactions, a two-way classification model was fitted for two loci at a time. When such digenic epistasis proved statistically significant, adjusted means were compared for each locus separately. When the interaction was non-significant, the interaction term was ignored and marginal means were computed for each locus.

Power of all tests was negatively affected by unequal cell sized in the factorial classification, which stems from the data structure of the population studied. However, the validity of a parametric test does not rest on a minimal sample size. As the provided data met the normality assumption, the tests are valid in small samples.

Results

Phenotypic variation

Flowering time ranged from 58.7 and 81.7 das with a mean of 70 das (Table 1). The EU accessions were predominantly late flowering, on average four to seven days after those from the remaining regions. Six-rowed accessions flowered on average about five days before two-rowed ones, but for both subgroups the range in flowering time was about 21 days. Both the genetic and genotype × environment interaction variances were highly significant (< 0.01), with former about eight fold larger than the latter. As a result, heritability was high (0.96).
Table 1

Means, minimum (min) and maximum (max) for flowering time (days after sowing, das) of all accessions, and of subsamples grouped by geographic origin and ear type

Germplasm

Number

Mean ± sd

Min

Max

Overall

220

70.0 ± 5.26

58.7

81.7

EU

108

72.9 ± 3.57

60.7

81.7

EA

39

65.8 ± 5.22

58.7

76.8

WA

44

68.1 ± 5.21

59.4

78.8

AM

29

68.1 ± 4.76

58.9

75.7

Two-rowed

126

72.1 ± 4.40

58.9

81.7

Six-rowed

94

67.2 ± 4.97

58.7

81.0

sd standard deviation, EU Europe, EA East Asia, WA West Asia and North Africa, AM America

Population structure

Using the mixed model, region of origin and ear row number explained, respectively, 26.1 and 21.6% of the genetic variation for flowering time (Table 2). A total of 173 alleles was observed across the EST-SSR loci (mean 6.9 alleles per locus). Using either the EM or the STRUCTURE approach, a two subgroup structure attracted the highest probability (Supplemental Figure S1). The QEM matrix for the mixed model, as estimated by the EM algorithm enjoyed a comparable fit relative to the one derived by STRUCTURE (Supplemental Table S4). In the subsequent association studies, a value of 2 was taken for KEM. Since both kinship matrices did not generate an improved fit (AIC) (Supplemental Table S4), this component was not included in the analyses.
Table 2

Percentage of explained genetic variance, var(G), for flowering time based on different mixed models

Model

% var(G)

QEM2

19.4

Row number

21.6

Origin

29.2

QEM2 + row number

22.9

QEM2 + origin

32.1

QEM2 + origin*

32.3

QEM2 + row number + origin*

33.9

* Subdivision into two origins: EU and non-EU

The background structure only accounted for 19.4% of the genetic variation in flowering time, so that each subgroup maintained plenty of variation. The distribution of SSR alleles was in close accordance with the taxonomic subgroups, as defined by ear row number (Fig. 1). The model-based and the row number-based assignment were in agreement for the bulk (>88%) of the accessions. The models adjusting for row number and QEM2 + row number explained almost the same proportion of the variation. In view of the distinct phenotypic differences between the EU and non-EU (origin*) accessions, as well as between the row number types, we adjusted the model to QEM2 + row number + origin* for the purpose of the association test.
https://static-content.springer.com/image/art%3A10.1007%2Fs00122-008-0896-y/MediaObjects/122_2008_896_Fig1_HTML.gif
Fig. 1

Calculated population structure, based on genotype at 25 EST-SSR loci. The population was partitioned into two color-coded subpopulations. Each bar represents a single accession, and the colored segments within each bar reflect the proportional contribution of the two subpopulations to that accession. Accessions are ordered by ear type (two-rowed and six-rowed) and by origin. EU Europe, EA East Asia, WA West Asia and North Africa, AM America

Molecular diversity and LD

Within the Ppd-H1 amplicon, 12 variable base pair positions were identified (Table 4), equivalent to a mean rate of one polymorphic site per 102 bp. Single nucleotide polymorphisms (SNPs) were more frequent in the coding than in the non-coding region (Table 3). In the coding region, five of the nine SNPs resulted in a codon replacement. The frequency of the SNP alleles was mostly well balanced, only at the ps1515 site the minor allele had a frequency (MAF) of <0.05. Six haplotypes (PpdH11 through PpdH16) were identified, four of which were present at a frequency > 0.05 (Table 4). In the non-EU material, most of the haplotypes were present at a frequency > 0.05. However, almost 85% of the EU accessions were of haplotype Ppd-H11. This same proportion of WA and AM accessions carried three, and EA as many as five haplotypes. Ppd-H11, -H12 and -H13 were represented in all the geographical origins, but the frequency of Ppd-H12 was <0.05 in the EU material. LD between the SNPs was highly variable (mean r2 of 0.41, ≤ 0.001) (Fig. 2).
Table 3

Estimates of nucleotide diversity (pi) and haplotype diversity (hd) in coding and non-coding regions of the genes Ppd-H1, HvCO1, and HvFT1

Genomic region

L

L*1

S

pi x 10−3 (± sd)

H

hd (± sd)

Ppd-H1 (N = 205)

Total

1,230

 

12

2.45 (± 0.17)

6

0.59 (± 0.03)

Coding

676

 

9

2.81 (± 0.24)

6

0.59 (± 0.03)

Non-coding

554

 

3

2.00 (± 0.13)

5

0.59 (± 0.03)

HvCO1 (N = 200)

Total

1,772

1771

16

1.55 (± 0.13)

10

0.68 (± 0.03)

Coding

911

 

5

0.20 (± 0.06)

5

0.12 (± 0.03)

Non-coding

861

860

11

2.98 (± 0.25)

7

0.66 (± 0.03)

HvFT1 (N = 195)

Total

1,975

1969

25

4.61 (± 0.07)

8

0.68 (± 0.02)

Total*2

972

 

8

1.51 (± 0.12)

6

0.66 (± 0.02)

Coding

437

 

1

0.12 (± 0.05)

2

0.05 (± 0.02)

Non-coding

535

 

7

2.67 (± 0.19)

6

0.66 (± 0.02)

5′-flanking

1,003

997

17

7.56 (± 0.06)

6

0.67 (± 0.02)

L fragment length in bp, L*1 fragment length after treating indels as single sites, S number of polymorphic sites, H haplotype number, sd standard derivation, N number of considered genotypes, total*2 5′-flanking region excluded

Table 4

Polymorphic sites in Ppd-H1 located in the exonic (e) and intronic (i) regions reflect their position in the AY970701 reference sequence

Haplotypes

Freq.

Polymorphic site position (ps)

1

4

4

3

1

5

1

5

1

7

2

5

1

7

8

5

1

8

3

9

1

8

4

3

2

0

3

2

2

0

3

5

2

0

3

6

2

2

3

0

2

3

2

2

2

3

2

7

i4

e5

i5

e6

e6

e6

e6

e6

e6

i6

e7

e7

   

s

s

s

  

s

 

s

 

AY970701

 

C

C

T

A

C

A

A

G

C

C

T

C

Ppd-H11

0.615

A

.

.

.

.

C

G

.

T

.

.

.

Ppd-H12

0.141

.

.

C

.

.

C

.

.

.

T

.

.

Ppd-H13

0.137

.

.

.

.

.

.

.

.

.

.

.

.

Ppd-H14

0.058

.

.

.

G

T

C

.

A

.

T

C

T

Ppd-H15

0.034

.

T

C

.

.

C

.

.

.

.

.

.

Ppd-H16

0.015

.

.

.

G

.

C

.

.

.

T

C

T

Model:QEM2

Associated

 

***

NS

***

**

*

**

***

*

***

***

**

**

var(G) explained

 

30.7

 

8.4

5.0

2.0

4.4

30.7

2.0

30.7

16.0

5.0

5.0

Model:QEM2 + row number + origin*

Associated

 

***

NS

*

*

NS

*

***

NS

***

***

*

*

var(G) explained

 

20.1

 

5.7

6.7

 

6.4

20.1

 

20.1

11.0

6.7

6.7

Replacement nucleotides are labeled “s”. Dots indicate identity with the reference sequence. Haplotype frequencies within the entire collection are shown. In the lower part of the figure, association tests between the polymorphic site and flowering time calculated with two models are summarized (NS = non-significant, * = ≤ 0.05, ** = ≤ 0.01, *** = ≤ 0.001). The % genetic variation explained, var(G) is shown for each significantly associated site

https://static-content.springer.com/image/art%3A10.1007%2Fs00122-008-0896-y/MediaObjects/122_2008_896_Fig2_HTML.gif
Fig. 2

Strength and extent of LD at Ppd-H1, HvCO1, and HvFT1. Polymorphic sites with a minor allele frequency of >0.01 were considered for pairwise calculation. Each point in the LD matrix represents a comparison between a pair of polymorphic sites, with the r2 values displayed below the diagonal, and P values for Fisher’s Exact test above. Points on the diagonal correspond to comparisons of each site with itself. Black, white and hatched margins indicate, respectively, exonic, intronic and 5′-flanking regions. Color codes for r2 and P values are given

Within HvCO1, 16 polymorphic sites (one polymorphic site per 111 bp) were defined, comprising 14 SNPs and two indels (Table 5). The non-coding sequence showed a moderate level of nucleotide diversity, but within the coding regions, this was reduced by 15 fold (Table 3). All five coding SNPs and two of the 11 non-coding polymorphic sites were present at a MAF of <0.05. Two (ps427 and ps582) of the five coding region sites defined a change in amino acid residue. Ten haplotypes (HvCO11 through HvCO110) present in at least two accessions were recognized, but six of these were rare (<0.05) (Fig. 3). Haplotype frequencies varied markedly between the geographical origins. HvCO11 was present in >75% of the EU accessions, while the other groups had a more balanced distribution, with the commonest haplotype present in no more than 40% of the accessions. Pairwise LD estimates produced a mean r2 value of 0.49 (≤ 0.001), with 88% of all highly significant associations (r2 > 0.5) located in intronic sequence (Fig. 2).
Table 5

Polymorphic sites in HvCO1 located in the exonic (e) and intronic (i) regions reflect their position in the AF490467 reference sequence

Haplotypes

Freq.

Polymorphic site position (ps)

1

8

2

4

2

7

5

0

6

5

8

2

8

4

3

8

8

5

9

1

2

1

0

9

3

1

2

6

7

1

3

4

8

1

4

5

5

1

4

8

9

1

5

7

6

1

5

9

6

1

8

1

6

2

0

0

8

e1

e1

e1

e1

i1

i1

i1

i1

i1

i1

i1

i1

i1

i1

i1

e2

 

s

 

s

            

AF490467

 

C

G

C

C

AT

A

C

A

A

A

A

T

C

C

T

G

HvCO11

0.520

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

HvCO12

0.129

.

.

.

.

.

.

G

.

G

.

C

C

A

T

C

.

HvCO13

0.129

.

.

.

.

.

del

.

.

.

.

.

.

.

.

C

.

HvCO14

0.114

.

.

.

.

del

.

.

.

.

.

.

.

.

.

C

.

HvCO15

0.030

.

.

.

.

.

.

.

G

.

.

.

.

.

.

C

.

HvCO16

0.020

.

.

.

T

.

del

.

.

.

.

.

.

.

.

C

.

HvCO17

0.020

.

.

T

.

.

.

G

.

G

G

C

C

A

.

C

A

HvCO18

0.020

.

.

.

.

.

.

.

.

.

.

.

.

.

.

C

.

HvCO19

0.010

T

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

HvCO110

0.010

.

A

.

T

.

del

.

.

.

.

.

.

.

.

C

.

Model: QEM2

Associated

 

NS

NS

*

*

NS

NS

**

NS

**

NS

**

**

**

NS

**

*

var(G) explained

   

2.1

1.2

  

2.7

 

2.7

 

2.7

2.7

2.7

 

4.1

2.1

Model: QEM2 + row number + origin*

Associated

 

NS

NS

NS

*

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

var(G) explained

    

1.2

            

Replacement nucleotides are labeled “s”. Dots indicate identity with the reference sequence. The starting point of indels is indicated. Haplotype frequencies within the entire collection are shown. In the lower part of the figure, association tests between the polymorphic site and flowering time calculated with two models are summarized (NS = non-significant, * = ≤ 0.05, ** = ≤ 0.01, *** = ≤ 0.001). The % genetic variation explained, var(G) is shown for each significantly associated site

https://static-content.springer.com/image/art%3A10.1007%2Fs00122-008-0896-y/MediaObjects/122_2008_896_Fig3_HTML.gif
Fig. 3

Haplotype frequency and distribution at Ppd-H1, HvCO1, and HvFT1 both across the whole collection and separately for geographical origin (EU Europe, EA East Asia, WA West Asia and North Africa, AM America). The number of relevant accessions is shown in brackets. Each hatched pattern represents a haplotype occurring at a frequency of >0.05. Rarer haplotypes are pooled, and represented by a white bar

Within HvFT1, 21 SNPs and four indels (one polymorphic site per 79 bp) were identified (Table 6). Among the eight haplotypes (HvFT11 through HvFT18) identified, only three were present at >0.05 (Fig. 3; Table 6). Across all the whole genotype set, the highest level of nucleotide diversity was in the 5′-flanking interval. The mean diversity in the intron regions was moderate, while the exon sequence revealed only a single SNP (Table 3).
Table 6

Polymorphic sites of the HvFT1 gene located in the exonic (e) and intronic (i) and 5′-flanking (5′) regions reflect their position in the DQ100372 reference sequence

Haplotypes

Freq.

Polymorphic site position (ps)

4

5

0

4

6

0

4

6

4

6

0

6

6

3

2

7

2

3

7

7

9

8

3

8

8

9

0

9

3

9

9

9

0

1

0

0

2

1

0

7

9

1

1

6

2

1

1

8

6

1

1

9

4

1

1

9

9

1

5

7

2

1

5

9

9

1

6

1

6

1

7

5

6

1

8

0

5

1

9

1

9

2

0

7

9

2

1

6

3

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

5′

i1

i1

i1

i1

i1

i1

i2

e3

DQ100327

 

C

C

A

C

G

G

del

G

A

C

G

G

T

CTTG

T

C

C

T

C

C

T

A

G

G

G

HvFT11

0.479

.

.

.

T

C

C

CGCG

A

G

G

del

C

C

del

C

G

T

.

T

.

.

T

C

.

.

HvFT12

0.273

G

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

T

C

.

.

HvFT13

0.124

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

HvFT14

0.036

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

del

.

.

.

.

.

.

.

HvFT15

0.031

.

.

.

.

.

C

CGCG

A

G

G

.

.

C

del

C

G

T

.

.

.

C

.

.

.

.

HvFT16

0.026

.

G

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

T

.

T

.

A

T

HvFT17

0.021

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

T

C

.

.

HvFT18

0.010

G

.

G

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

T

C

.

.

Model: QEM2

Associated

 

***

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

***

***

NS

NS

var(G) explained

 

6.6

                    

8.8

9.7

  

Model: QEM2 + row number + origin*

Associated

 

*

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

NS

***

***

NS

NS

var(G) explained

 

3.2

                    

6.0

7.2

  

Dots indicate identity with the reference sequence. The starting point of indels is indicated. Haplotype frequencies within the entire collection are shown. In the lower part of the figure, association tests between the polymorphic site and flowering time calculated with two models are summarized (NS = non-significant, * = ≤ 0.05, ** = ≤ 0.01, *** = ≤ 0.001). The % genetic variation explained, var(G) is shown for each significantly associated site

A strong LD block (ps606–ps1199) was detected in the 5′-flanking region, comprising at least 590 bp and flanked by sequence with a highly variable pairwise LD structure (Fig. 2). This interval had a mean r2 of 0.88 (≤ 0.001), while the mean LD declined sharply to 0.40 (≤ 0.001) in the coding region. This LD block structure was attributable to a split into two tight haplogroups—haplogroup I comprising haplotype HvFT11 and HvFT15, and haplogroup II with HvFT12 through HvFT14, and HvFT16.

Interlocus associations were minor compared to intralocus LD, but some differences were apparent between the locus pairs (Supplemental Figure S2). The mean level of significant LD between Ppd-H1HvFT1 (r2 = 0.05; ≤ 0.05, MAF > 0.01) was twofold less than that observed for Ppd-H1HvCO1 (r2 = 0.11; ≤ 0.05, MAF > 0.01) and HvCO1HvFT1 (r2 = 0.10; ≤ 0.05, MAF > 0.01).

Haplotype–phenotype association

Assuming a population structure (defined by KEM = 2), significant haplotype/flowering time associations were identified for all three genes (Table 7). The Ppd-H1 association explained 32.4% of the genetic variance, and the haplotype classes revealed highly significant differences of five to ten days between accessions carrying the latest flowering Ppd-H11 haplotype and those carrying any of the five remaining ones (Table 8). One replacement (ps2036), one silent (ps2032) and three intronic sites (ps1725, ps1443, ps2230) were highly significantly (< 0.001) associated with phenotype. The three sites, ps1443, ps2032, ps2036, were in perfect LD to one another (Fig. 2) and are diagnostic for Ppd-H11; these explain 30.7% of the genetic variation (Table 4).
Table 7

Estimated percentage of genetic variance, var(G), for flowering time explained by haplotypes of the genes Ppd-H1, HvCO1, and HvFT1 based on two mixed models

Models for SNP genotypes

Model QEM2

Model QEM2 + row number + origin*

% var(G)

P value#

% var(G)

P value#

Ppd-H1

32.4

<0.0001

22.4

<0.0001

HvCO1

9.3

0.0005

0

0.1112

HvFT1

9.5

0.0003

3.4

0.0621

Ppd-H1 + HvCO1 + Ppd-H1 × HvCO1

51.0

<0.0001

48.8

<0.0001

Ppd-H1 + HvFT1 + Ppd-H1 × HvFT1

41.5

0.6544

30.6

0.8584

HvCO1 + HvFT1 + HvCO1 × HvFT1

16.9

0.0450

12.3

0.0298

* Subdivision into two origins, EU and non-EU

Significance level of Wald-t test; in the case of the 2-factorial models P values refer to the interaction

Table 8

Means and their 95% confidence intervals of the haplotype classes of Ppd-H1, HvCO1 and HvFT1 for flowering time based on two models

Haplotypes

Model

Model

QEM2

QEM2 + row number + origin*

Mean

CI95

P§

 

Mean

CI95

 

P§

Ppd-H11

72.56

0.73

.

b

72.06

0.86

.

b

Ppd-H12

66.20

1.50

a

.

67.05

1.54

a

.

Ppd-H13

67.38

1.47

a

.

67.85

1.46

a

.

Ppd-H14

65.67

2.54

a

.

66.56

2.52

a

.

Ppd-H15

66.88

3.21

a

.

67.80

3.18

a

b

Ppd-H16

61.66

4.53

a

.

62.47

4.42

a

.

HvCO11

71.61

1.15

.

b

70.85

1.29

a

.

HvCO12

68.21

1.94

a

b

69.25

1.90

a

.

HvCO13

71.25

2.13

.

b

70.20

2.12

a

.

HvCO14

67.09

2.22

a

b

68.33

2.14

a

.

HvCO15

67.83

3.81

a

b

68.99

3.58

a

.

HvCO16

63.66

4.23

a

.

64.67

3.98

a

.

HvCO17

63.08

5.30

a

b

64.31

4.98

a

.

HvCO18

70.67

4.49

a

b

71.41

4.34

a

.

HvCO19

70.14

6.33

a

b

68.64

6.10

a

.

HvCO110

68.23

6.35

a

b

67.23

5.94

a

.

HvFT11

70.50

0.99

a

b

69.82

1.00

a

.

HvFT12

68.40

1.33

a

.

68.99

1.26

a

.

HvFT13

73.70

1.95

.

b

72.39

1.92

a

.

HvFT14

71.61

3.38

a

b

70.09

3.30

a

.

HvFT15

72.53

3.99

a

b

72.92

3.92

a

.

HvFT16

71.61

4.12

a

b

71.57

3.86

a

.

HvFT17

65.31

4.50

a

.

67.32

4.42

a

.

HvFT18

70.09

6.34

a

b

68.85

5.96

a

.

Adjusted for PpdH1

 HvFT11

67.37

1.42

a

.

67.48

1.40

a

.

 HvFT12

65.93

1.30

a

.

66.19

1.40

a

.

 HvFT13

66.26

2.46

a

.

65.76

2.56

a

.

 HvFT14

65.88

3.11

a

.

66.34

3.08

a

.

 HvFT15

60.61

5.25

a

.

61.37

5.22

a

.

 HvFT16

69.33

3.89

a

.

68.93

3.88

a

.

 HvFT17

67.47

3.89

a

.

68.83

3.96

a

.

 HvFT18

62.60

5.43

a

.

62.71

5.36

a

.

Adjusted for HvFT1

 Ppd-H11

71.76

1.28

.

b

72.00

1.38

.

b

 Ppd-H12

63.40

2.14

a

.

63.67

2.24

a

.

 Ppd-H13

65.49

1.83

a

.

65.99

1.86

a

.

 Ppd-H14

65.11

3.00

a

.

65.43

3.00

a

.

 Ppd-H15

67.24

3.70

a

b

67.36

3.70

a

b

 Ppd-H16

61.09

4.58

a

.

61.25

4.54

a

.

§ Means followed by a common letter are not significantly different according to the Bonferroni-Holm procedure

In contrast, the HvCO1 and HvFT1 haplotypes were not able to differentiate between the early and the late-flowering types, as, respectively, 80 and 62% of the haplotype classes were associated with an intermediate phenotype (Table 8). At HvCO1, only intronic sites showed any significant (≤ 0.01) association with flowering time (Table 5). Five of these, all in perfect LD with one another (ps912, ps1267, ps1455, ps1489, ps1576), explained 2.7%, while ps1816 explained 4.1% of the genetic variation (Fig. 2, Table 5). At HvFT1 the 5′-flanking region SNP at ps450 and two intronic sites (ps1805 and ps1919) in high LD with one another (r2 = 0.8; ≤ 0.0001) were significantly (P ≤ 0.001) associated with flowering time, but explained, respectively, only 6.6, 8.8, and 9.7% of the genetic variation (Fig. 2; Table 6).

Digenic epistasic interaction was highly significant for the Ppd-H1/HvCO1 combination, weakly so for HvCO1/HvFT1 and non-significant for Ppd-H1/HvFT1 (Table 7). As Ppd-H1 exerted the largest influence on flowering time, the impact of allelic distribution at this gene on the association results for HvFT1 and HvCO1 was evaluated. Excluding the non-significant gene × gene interaction from the model, the fixed HvFT1 effects were calculated by adjusting for the effect of Ppd-H1, leading to a dramatic reduction in the significance of the association between HvFT1 and flowering time. Phenotypic means of the various haplotype classes did not differ significantly from one another (Table 8). In contrast, a similar adjustment for the effect of HvFT1 on Ppd-H1 did not affect the differentiation between the late flowering PpdH11 haplotype and the remaining haplotypes (Table 8). Given the statistically significant Ppd-H1 × HvCO1 interaction, the Ppd-H1 background was fixed for Ppd-H11 and the earlier flowering haplotypes PpdH12–6 to estimate the effect of HvCO1 within these haplotype classes (Fig. 4a). One group of haplotypes (HvCO11, HvCO12, HvCO16) suggested a background-dependent change in phenotypic expression, while a second group (HvCO13, HvCO14, HvCO15, HvCO110) behaved almost independently. In the presence of Ppd-H11, HvCO11 and HvCO12 generated the latest flowering phenotype, differing significantly (by 8 days) from carriers of HvCO14. This earlier flowering haplotype can be identified by a two-base pair deletion (site 843; Table 5). For all other haplotyes resulting in a significantly different phenotype it was not feasible to generate a diagnostic marker. In the PpdH12–6 background the flowering time of HvCO11 and HvCO12 carriers was accelerated to become indistinguishable from carriers of HvCO14.
https://static-content.springer.com/image/art%3A10.1007%2Fs00122-008-0896-y/MediaObjects/122_2008_896_Fig4_HTML.gif
Fig. 4

The interaction between Ppd-H1 and HvCO1. The estimated mean phenotypic values are shown for HvCO1 haplotypes in carriers of the PpdH11 and Ppd-H12−6 haplotypes, applying a the model QEM2, and bQEM2 + row number + origin*. Haplotypes are denoted by exponents attached to the gene symbol. Means followed by a common letter do not differ significantly from one another, using the Bonferroni–Holm procedure

For HvCO1/HvFT1 the haplotype of each gene was changed in a stepwise manner. In a HvCO11 background, significant differences between the HvFT1 haplotype classes were noted, whereas in the HvCO12–10 backgrounds no such significant differences occurred. No HvFT1 haplotype background was associated with any significant variation in the presence of a particular set of HvCO1 haplotypes (data not shown).

Finally, epistatic interactions were sought using the model QEM2 + row number + origin* which explained most of the phenotypic variation in flowering time (Table 2). Only Ppd-H1 was consistently associated with flowering time (Table 7). HvCO1 was not associated with flowering time, although the interaction between Ppd-H1 and HvCO1 was highly significant. Once again, significant phenotypic differences were observed between the HvCO1 haplotype classes in the presence of Ppd-H11, but none in the Ppd-H12–6 background (Fig. 4b).

Discussion

The genetic dissection of quantitative variation is key to the implementation of genomics-based crop improvement strategies. Association mapping can be a powerful means of understanding the genetic basis of quantitative variation, and can even succeed in identifying candidate genes involved in its determination (review: Hirschhorn et al. 2002). We have investigated here the allelic effects of three genes involved in the determination of flowering time, and have used a haplotype-based association analysis in conjunction with statistical models of differing complexity. A special focus was on the impact of gene × gene interactions on the outcome of association results.

Population structure, the target genomic region, and the number of polymorphic sites can all have a major impact on the pattern of LD and consequently, on the outcome of association studies (Akey et al. 2003; Ke et al. 2004; Pritchard et al. 2000b). The population structure of the barley collection is largely consistent with the phenotype-based grouping into two- and six-rowed ear types, as has been noted in other sets of spring (Brantestam et al. 2007; Hayes and Szucs 2006) and winter cultivars (Matus and Hayes 2002; Stracke et al. 2007; Thiel et al. 2003). Ear type is encoded by a single major gene v, located on chromosome 2H (Komatsuda et al. 2007), which cannot account for the existence of genetically distinct subpopulations. However, breeders tend to avoid intermixing these types, as six-rowed barleys are usually destined for feed, and the two-rowed ones for malting (Hayes and Szucs 2006; Kjaer and Jensen 1996; Matus and Hayes 2002).

While the level of nucleotide diversity in the non-coding regions of the three genes investigated was comparable, coding sequences revealed marked differences. The coding region of Ppd-H1 is particularly diverse, with a substantial number of replacement alleles. This is consistent with the reported role of the A. thaliana pseudo-response regulator gene family (PRR), where both PRR7 (orthologous to Ppd-H1) and PRR5 are rich in allelic variation allowing for the wide adaptation of the species (Michael et al. 2003). The late flowering haplotype Ppd-H11 is synonymous with the ppd-H1 allele present in the cultivar Triumph (Turner et al. 2005). Surprisingly, in addition to the diagnostic SNPs for Ppd-H11, two polymorphic intronic sites were also significantly associated with flowering time, but these are independent of the dimorphism between the late and the earlier flowering haplotypes. Given that the assignment to subgroups only imperfectly reflects the genetic background of the collection (Balding 2006), this further association may be attributable to residual background structure within the subgroups.

Many rare exonic SNPs are present in HvCO1, and this resulted in the definition of many haplotypes and a low average nucleotide diversity. A. thalianaCO belongs to a family of putative transcription factors characterized by two conserved domains. B-boxes near the N-terminus regulate protein-protein interactions, and a CCT domain near the C-terminus controls the nuclear localization of the CO protein (Robson et al. 2001) and binds to a trimeric CCAAT factor (Ben-Naim et al. 2006; Robson et al. 2001). All of the functional CO mutations reported to date are located in one or other of these two domains, which underlines their importance to the control of flowering time (Ben-Naim et al. 2006; Lagercrantz and Axelsson 2000; Robson et al. 2001). In HvCO1 one polymorphic non-replacement site was identified in each of the two domains. As resequencing was incomplete (76% coverage), it is possible that a number of other polymorphic sites was not identified. Nevertheless, because of LD structure, we assume that all possible associations have been identified.

HvFT1 has a divergent diversity structure both up- and downstream of the start codon. The extended LD block in the 5′-flanking region has been noted previously in both spring and winter barley cultivars (Yan et al. 2006). As this feature is associated neither with flowering time nor with growth habit, it may either play a role in some other signaling pathway, or be an artifact of genetic drift. In A. thaliana, the FT promoter region is thought to be a potential target for selection for early flowering (Roux et al. 2006) and FT is one of a set of genes which is regulated by photoperiodic, vernalization, gibberellin, and developmental signals (Abe et al. 2005; Ausin et al. 2005; Kardailsky et al. 1999; Wigge et al. 2005). Induced mutations at FT have revealed the importance of binding sites at the promoter (Helliwell et al. 2006; Huang et al. 1995), the first intron (Helliwell et al. 2006) and of potential binding pockets in the coding sequence (Hanzawa et al. 2005). A motif search in HvFT1 revealed that ps460 and ps464 in the 5′-flanking region are both located at a potential binding site of a MADS-box transcription factor. However, the minor allele generates a change in the potential binding motif at both sites, but occurred at a frequency of <0.05 and neither site was associated with flowering time. Interestingly, the two intron SNPs that have been recently proposed to be diagnostic between spring and winter barleys (Yan et al. 2006; Vrn-H3, SNP270 and SNP384) are identical to the segregating sites SNP1805 and SNP1819 in our spring barley collection. As shown in our study, the potential effect on flowering time of both SNPs loses on significance when adjusting for Ppd-H1. These contradictory results show that sampling effects (number of genotypes and genetic background) have a strong influence on the results of association studies.

Overall, our data demonstrate that most of the variation in flowering time in the collection is not caused by functional mutations in either HvCO1 or HvFT1. This seems to be consistent with QTL results from the literature. The location of HvCO1 did not coincide with the maximum-likelihood peak for flowering QTL (Griffiths et al. 2003), and though HvFT1 was located in a QTL for the earliness factor eps7S, no polymorphism between the cross parents Igri and Triumph was detected (Faure et al. 2007). However, it is still possible that induced mutations in the putative functional domains of both genes could show an analogous effect on flowering time in barley as they do in A. thaliana.

The phenotypic differences between the HvFT1 haplotype classes were mainly attributable to an imbalanced distribution of Ppd-H1 alleles (Supplemental Table S5). The proportion of the late-flowering Ppd-H11 carriers in the three most frequent HvFT1 haplotype classes was 70.6, 35.8 and 95.8% for, respectively, HvFT11, HvFT12 and HvFT13, which led to the identification of spurious significant differences among the haplotype classes. Although HvFT1 is a key gene for the control of flowering time (Turner et al. 2005), none of the variants exerted any effect when the data were corrected for the effect of Ppd-H1. Direct biological inferences from statistical tests are notoriously risky (Cordell 2002), but our conclusions are strongly supported by the lack of difference in the expression level of Ppd-H1 between cultivar Triumph (Ppd-H11) and a closely related line carrying the early flowering Ppd-H13 allele of cultivar Igri (Turner et al. 2005). However, the expression level of HvFT1 was strongly dependent on the identity of the allele at Ppd-H1. HvCO1 expression was high in the presence of both Ppd-H1 alleles, but was significantly lower in the Ppd-H11 background at some measurement time points. As the contrasting genotypes were monomorphic at HvCO1, biological evidence for the Ppd-H1 × HvCO1 interaction remains to be collected. It is of interest, however, that in rice, epistatic interactions affect all combinations of Hd1 (HvCO1), Hd2 (Ppd-H1) and Hd3a (HvFT1) (Lin et al. 2000).

As shown in our study, genotypes with identical Ppd-H1 background revealed significant differences in the flowering time depending on the allelic constitution at HvCO1. Hence, it might be useful for practical breeding purposes to select not only for Ppd-H1 but also for HvCO1 alleles. With exception of HvCO14, however, it was not feasible to generate diagnostic markers for the sequences at HvCO1 in our data set. Thus, to date re-sequencing might be the only one, but cost-intensive method for the identification of the desirable haplotype.

It has not yet been possible to decide, whether the highly significant Ppd-H1 × HvCO1 interaction is a result of co-evolution, or whether it is due to a sampling artifact. Interlocus LD was significant, but rather weak (mean values of r2 = 0.05 and 0.11). The imbalanced distribution of the alleles in the EU and non-EU (EA, WA, AM) materials may reflect a degree of local adaptation (Supplemental Table S6). Late flowering favors vegetative growth, and thus enhances the amount of assimilate accumulated by the plant where moisture and temperature remain non-limiting; in contrast, early flowering is advantageous where terminal drought or temperature stress is commonplace (Roux et al. 2006). The strikingly high frequencies of the Ppd-H11 and HvCO11 haplotypes across the whole collection (76% of the accessions are Ppd-H11/HvCO11) is mainly due to their predominance within the EU set, which contributed 76.5% of all entries with this combination. The effect of narrow domestication-related bottlenecks on haplotype diversity between modern barley cultivars and its closest wild relative H.spontaneum has been described repeatedly (Badr et al. 2000; Matus and Hayes 2002; Piffanelli et al. 2004).

Consequences for association studies

The current study illustrates the potential of association mapping but also highlights some common pitfalls. The outcomes agree well with a QTL-based analysis in which variation at Ppd-H1 accounted for some 60% of the genetic variation in flowering time (Laurie et al. 2004). Note that the association mapping approach delivered a much lower estimate for this proportion. This might be partly attributable to the fact that in the QTL approach only two alleles of each locus were considered from parent lines strongly contrasting in flowering time, whereas our association approach was based on a wide range of germplasm with different allelic compositions.

Results from the present study and from literature (Laurie 1997; Laurie et al. 1994, 2004) indicated both, QTL analysis and association approach, are suitable tools for mapping of quantitative loci with strong effects on flowering time. For future research gene identification might greatly facilitate by the recently suggested integrative method nested association mapping (Yu et al. 2008) where both approaches are combined by using first QTL mapping in sets of multi-line mapping populations to detect chromosomal regions of interest and second, association mapping with high-density markers for fine dissection. Traits controlled by polygenic networks involving a large number of small-effect QTL are probably not readily amenable to the association approach. A recent theoretical treatment that has highlighted the impact of epistasis on QTL detection in association studies was provided by Jannink (2007). Our analysis also underlines the need to consider gene × gene interactions, both to highlight spurious associations and to identify hidden variation.

Acknowledgments

The field trials were supported by the breeding companies Lochow Petkus GmbH (Bergen-Wohlde) and Dr. J. Ackermann & Co (Irlbach). The authors thank K. Trnka, J. Jesse, S. Koch and A. Smyra for their excellent technical assistance, and M. Mohr for her valuable help. This work was financially supported by the German Federal Ministry of Education, and Research (grants 0313098A-C).

Open Access

This article is distributed under the terms of the Creative Commons Attribution Noncommercial License which permits any noncommercial use, distribution, and reproduction in any medium, provided the original author(s) and source are credited.

Supplementary material

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Copyright information

© The Author(s) 2008

Authors and Affiliations

  • Silke Stracke
    • 1
    • 2
  • Grit Haseneyer
    • 1
    • 3
  • Jean-Baptiste Veyrieras
    • 4
  • Hartwig H. Geiger
    • 5
  • Sascha Sauer
    • 6
  • Andreas Graner
    • 1
  • Hans-Peter Piepho
    • 7
  1. 1.Leibniz Institute of Plant Genetics and Crop Plant Research (IPK)GaterslebenGermany
  2. 2.Department of Crop Sciences, Quality of Plant ProductsUniversity of GöttingenGöttingenGermany
  3. 3.Plant BreedingTechnische Universität München/Centre of Life and Food Sciences WeihenstephanFreisingGermany
  4. 4.UMR, INRA UPS-XI INAPG CNRS, Génétique VégétaleGif-sur-YvetteFrance
  5. 5.Institute of Plant Breeding, Seed Science, and Population GeneticsUniversity of HohenheimStuttgartGermany
  6. 6.Max-Planck-Institute for Molecular GeneticsBerlin-DahlemGermany
  7. 7.Institute for Crop Production and Grassland ResearchUniversity of HohenheimStuttgartGermany