Phylogenetic reconstruction at the species and intraspecies levels in the genus Pisum (L.) (peas) using a histone H1 gene

Gene 504 (2012) 192–202

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Phylogenetic reconstruction at the species and intraspecies levels in the genus Pisum (L.) (peas) using a histone H1 gene Olga O. Zaytseva a, b, Vera S. Bogdanova a, Oleg E. Kosterin a, b,⁎ a b

Institute of Cytology and Genetics SB RAS, Acad. Lavrentyev ave. 10, Novosibirsk, 630090, Russia Novosibirsk State University, Pirogova str. 2, Novosibirsk, 630090, Russia

a r t i c l e

i n f o

Article history: Accepted 12 May 2012 Available online 18 May 2012 Keywords: Histone H1 Pisum L. Phylogenetic trees PsbA-trnH chloroplast intergenic spacer Phylogenetic marker

a b s t r a c t A phylogenetic analysis of the genus Pisum (peas), embracing diverse wild and cultivated forms, which evoke problems with species delimitation, was carried out based on a gene coding for histone H1, a protein that has a long and variable functional C-terminal domain. Phylogenetic trees were reconstructed on the basis of the coding sequence of the gene His5 of H1 subtype 5 in 65 pea accessions. Early separation of a clear-cut wild species Pisum fulvum is well supported, while cultivated species Pisum abyssinicum appears as a small branch within Pisum sativum. Another robust branch within P. sativum includes some wild and almost all cultivated representatives of P. sativum. Other wild representatives form diverse but rather subtle branches. In a subset of accessions, PsbA-trnH chloroplast intergenic spacer was also analysed and found less informative than His5. A number of accessions of cultivated peas from remote regions have a His5 allele of identical sequence, encoding an electrophoretically slow protein product, which earlier attracted attention as likely positively selected in harsh climate conditions. In PsbA-trnH, a 8 bp deletion was found, which marks cultivated representatives of P. sativum. © 2012 Elsevier B.V. All rights reserved.

1. Introduction Histone H1 is among the major protein components of eukaryotic chromatin. It participates in chromatin condensation into higher-order structure by binding linker DNA (Allan et al., 1986; Bharath et al., 2003). Histone H1 undergoes an intensive turnover in chromatin (Bustin et al., 2005; Misteli et al., 2000; Th'ng et al., 2005; Zlatanova et al., 2000), its relative abundance varies among tissues (Fan et al., 2003, 2005; Pearson et al., 1984) and correlates with the nucleosome repeat length (Routh et al., 2008; Woodcock et al., 2006), so H1 is a flexible component of ‘molecular environment’ for transcription machinery. In contrast to the core histones, histone H1 is a highly variable protein (Doenecke et al., 1997; Happel and Doenecke, 2009) with intraspecific polymorphism observed in many unrelated groups (Kosterin et al., 1994). In most organisms there are several non-allelic subtypes of histone H1 which may have some functional peculiarities (Cole, 1987; Happel and Abbreviations: Cox1, cytochrome c oxidase subunit I gene; His(2–6), group of linked pea genes coding for histone H1 subtypes 2–6; His-3, pea histone H1 subtype 3 gene; His5, pea histone H1 subtype 5 gene; ICG, Institute of Cytology and Genetics; IChBFM, Institute of Chemical Biology and Fundamental Medicine; SB RAS, Siberian Branch of Russian Academy of Sciences; ML, maximum likelihood; MP, maximum parsimony; NJ, Neighbour-Joining; PsbA, core protein of photosystem II gene; RbcL, large subunit of ribulose-1,5-bisphosphate carboxylase oxygenase gene; SCA, seed cotyledon albumin gene; SD, standard deviation; trnH, histidine tRNA gene; PsH1b, Pisum sativum histone H1 b gene. ⁎ Corresponding author at: Institute of Cytology and Genetics SB RAS, Acad. Lavrentyev ave. 10, Novosibirsk, 630090, Russia. Tel.: +7 383 363 49 36; fax: +7 383 333 12 78. E-mail address: [email protected] (O.E. Kosterin). 0378-1119/$ – see front matter © 2012 Elsevier B.V. All rights reserved. doi:10.1016/j.gene.2012.05.026

Doenecke, 2009; Izzo et al., 2008; Ponte et al., 1998; Sancho et al., 2008), such as their ability to bind chromatin, intensity of turnover in chromatin (De et al., 2002; Th'ng et al., 2005), effect on nucleosomal spacing (Oberg et al., 2012; Sancho et al., 2008). Some H1 subtypes may bring about differences in chromatin compaction (Alami et al., 2003; Clausell et al., 2009; Khadake and Rao, 1995; Liao and Cole, 1981; Talasz et al., 1998), have different content in transcriptionally active and inactive chromatine sites (Parseghian et al., 2000, 2001; Th'ng et al., 2005) and participate in regulation of expression of specific set of genes (Jedrusik and Schulze, 2001; Sancho et al., 2008; Takami et al., 2000). The H1 subtype spectrum usually changes through ontogenesis and in the process of cell differentiation (Cole, 1987; Doenecke et al., 1997; Ponte et al., 1998). The total histone H1 amount and relative content of different subtypes may be important factors in differential gene regulation and cell differentiation in ontogenesis (Brown et al., 1996, 1997; Sera and Wolffe, 1998; Th'ng et al., 2005). At the same time variation in the histone H1 molecule structure may participate in fine tuning of expression of many genes (Alami et al., 2003) and hence contribute to adaptive evolution (Berdnikov et al., 1993a,b). In insects, correlation was observed between the intra-order variation of H1 molecule length and the number of species in the order rather than with the order's evolutionary age (Berdnikov et al., 1993b). The study of geographic patterns of distribution of allelic variants of pea H1 subtypes revealed that one particular allele of subtype 5 was most abundant in the regions with low accumulated temperature of the vegetation period (Berdnikov et al., 1993a). Comparison of nearly isogenic pea lines revealed an effect of substitution of allelic variants of certain

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H1 subtypes on some quantitative traits (Berdnikov et al., 1999, 2003; Bogdanova et al., 1994, 2007). Because of their high variability, histone H1 genes could be helpful for phylogenetic reconstructions at intraspecific level. Moreover, frequent indels in the C-terminal domain may serve as unique markers of lineages (Trusov et al., 2004). One of the most suitable model organisms for investigation of linker histone variation influence on phenotype is the genus Pisum L. in which intraspecies polymorphism was found for all 7 unique paralogous histone H1 genes coding for different subtypes (Kosterin et al., 1994). The aim of the present study was to reconstruct phylogenetic relationships in the genus Pisum using nucleotide sequences of the gene His5 coding for H1 subtype 5 (Bogdanova et al., 2005). The taxonomy of the genus is complicated (Smýkal et al., 2010) due to great diversification of local populations, as evidenced, for instance, by highly diverse geno- and phenotypes originated from a small territory of Central Israel (Ben-Ze'ev and Zohary, 1973) and peas of different evolutionary lineages co-occurring in some provinces of Turkey (Kosterin et al., 2010). Pea is mostly a self-pollinator, therefore genetic exchange between populations through cross-pollination is limited, although still possible (Bogdanova and Berdnikov, 2000; Loenning, 1984). At present, two species are distinguished by most authors: Pisum sativum L., represented by diverse wild and cultivated forms and a clear-cut and reproductively isolated wild species Pisum fulvum Sibth. et Smith confined to arid regions of Anterior Asia (Ben-Ze'ev and Zohary, 1973; Makasheva, 1979). Most authors (e.g. Ellis et al., 1998; Kosterin and Bogdanova, 2008) also consider Pisum abyssinicum A. Br. as a separate species cultivated in South Arabia and Ethiopia. Both P. fulvum and P. abyssinicum are reproductively isolated from P. sativum, as their hybrids with the latter suffer from sterility due to some chromosomal rearrangements (Ben-Ze’ev and Zohary, 1973; Conicella and Errico, 1990) and genetic incompatibility (Bogdanova and Kosterin, 2007 and unpublished). Less severe reproductive barriers associated with nuclear-cytoplasm incompatibility (Bogdanova and Berdnikov, 2001; Bogdanova et al., 2009) and translocations (BenZe'ev and Zohary, 1973) exist as well within P. sativum, subdivided into two distinct karyotypic forms. Wild representatives of P. sativum were traditionally subdivided into two groups: tall mesophilous plants confined to Mediterranean maquis were attributed to P. sativum subsp. elatius (Bieb.) Schmal., while rather low xerophytic forms growing in arid zones of the Middle East — to ‘P. humile Boiss et Noe’ (its valid name is P. sativum subsp. syriacum Berger) (Ben-Ze'ev and Zohary, 1973; Makasheva, 1979). However, the results of karyological studies contradicted such division (Ben-Ze'ev and Zohary, 1973). Presently most authors following Townsend (1968) and Davis (1970) accept rather conventionally that all wild representatives of P. sativum belong to the subspecies P. sativum subsp. elatius (Bieb.) Schmahl. sensu lato, opposed to cultivated subspecies P. sativum subsp. sativum L. Several attempts were made recently to reconstruct phylogenetic relationships among peas (Ellis et al., 1998; Hoey et al., 1996; Jing et al., 2007, 2010; Lu et al., 1996; Vershinin et al., 2003). All of them resulted in P. fulvum and P. abyssinicum to form separate branches on the trees, but the branch of P. fulvum was basic while that of P. abyssinicum resided within P. sativum, making the latter a paraphyletic species. The position of different groups within P. sativum varied. For example, a reconstruction based on morphological, allozyme and RAPD markers (Hoey et al., 1996) was consistent with karyologic studies of the same lines (BenZe'ev and Zohary, 1973). On the phylogenetic trees obtained by Ellis et al. (1998) and Jing et al. (2007), topology of branches leading to accessions attributed to ‘P. humile’ was uncertain: some resided among cultivated P. sativum forms and one among ‘P. elatius’, proving the taxon ‘humile’ to be artificial. Earlier we investigated three functionally unrelated molecular markers from different cellular genomes: RbcL (plastid), CoxI (mitochondrial) and SCA (nuclear) in diverse pea accessions (Kosterin and Bogdanova, 2008; Kosterin et al., 2010), two allelic variants being

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found in each marker. Four of 8 possible allele combinations of 3 markers appeared frequently found and were designated A, B, C and D (other combinations referred to as ‘rare’). Their mutational transitions presumably followed the order A (RbcL+, Cox1+, SCAf) → C (RbcL +, Cox1−, SCAf) → D (RbcL−, Cox1 −, SCAf) → B (RbcL−, Cox1−, SCAs) (Kosterin et al., 2010). Almost all cultivated P. sativum represented combination B (very few had D), all four combinations were found among wild representatives of P. sativum while P. fulvum and P. abyssinicum had combination A. Three above mentioned mutations supposedly happened to mark some events in evolution of wild P. sativum, such as expansion of wild peas with combination A from Anterior Asia westwards, followed by almost a backward migration of carriers of the derivative combination B (Kosterin et al., 2010). Hence these four combinations will be used below as a kind of an informal intraspecies ‘operational taxonomy’ within P. sativum, the actual intraspecies taxonomy being still insufficiently elaborated. In this work we sequenced the His5 gene of histone H1 subtype 5 (for the gene nomenclature see Kosterin et al., 1994) of most wild and cultivated pea accessions involved in (Kosterin and Bogdanova, 2008; Kosterin et al., 2010) and constructed phylogenetic trees based on these sequences. In a smaller set of peas, we compared variation in His5 with that of PsbA-trnH chloroplast intergenic spacer frequently used in phylogenetic studies at the interspecific level and proposed as a marker for genetic barcoding in plants (Kress et al., 2005, Kress and Erickson, 2007). The His5 sequences appeared informative while PsbA-trnH not enough variable. 2. Materials and methods 2.1. Plant material Sixty five accessions of pea germplasm were analysed. Of them, 61 were characterised in detail in Table 1 in (Kosterin and Bogdanova, 2008) and Table 1 in (Kosterin et al., 2010). These represent the following here accepted taxa: P. fulvum: L95, WL2140, VIR2523, VIR3397, VIR6070, VIR6071, 701, 702, 703, 706, 707, 708; P. abyssinicum: JI1876, WL1446, VIR2759, VIR3567; P. sativum subsp. elatius (wild): CE1, CE2, JI254, JI1091, JI1092, JI1093, JI1094, JI1095, JI1096, JI1794, JI2055, JI2724, JI3553, L100 (=712), Ps002, Ps008, P012, P013, P015, P016, P017, PI343993, PI344008, PI344537, VIR320, VIR320*, VIR1851, VIR2521, VIR2524, VIR2998, VIR4014, VIR7327, WL805, WL2123, 711, 713, 714, 721, 722, 723; P. sativum subsp. sativum (cultivated) JI2105, Pa014; and two subspecies of dubious validity: P. sativum subsp. jomardii (Schrank) Kosterin from Egypt: VIR3424, VIR3439, and P. sativum subsp. transcaucasicum Govorov from Georgia: VIR3249. Besides, we added 4 accessions of P. sativum subsp. sativum: VIR2514, Syria, local form; VIR4362, Russia, Vologda Province, local form; VIR6560, Tajikistan, local form; WL1238, a popular testerline by Herbert Lamprecht, all, except for VIR2514 which was not analysed, with combination B of alleles of the three markers (see Introduction). Partial His5 sequences were also obtained from accessions VIR3913, VIR3971, VIR6135. These accessions were not analysed for SCA, RbcL and Cox1. 2.2. DNA isolation, PCR-amplification and sequencing DNA was isolated following (Bogdanova et al., 2009). Eight primers were designed on the basis of nucleotide sequence of VIR4362 His5 gene with adjacent non-coding regions (EMBL ID: AJ543403) (Table 1). These primers were used in different combinations to specifically amplify His5 but not other paralogous H1 genes. Primers derived from the coding region were used as additional ones for the Sanger reaction. Disposition of the primers is shown in a supplementary Fig. s1. The spacer PsbA-trnH was sequenced using primers PsbA3′f adopted from (Sass et al., 2007) and the originally designed trnH1r (Table 1).

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Table 1 Primers used to amplify and sequence the His5 gene in peas. Primer

Sequence 5′–3′

Matching sequence, position of the first nuceotide with respect to His5 coding position 1, direction

H1-514f H1-52f H1-591f H1-59f H1-53f cd7up2 H1-52r H1-54r H1-513r H1-55r H1-551r PsbA3′f trnH1r

CATCCCCATCCAAGGGCTGA CCACACTCATTTCACTATTTAAACC CATGAACCGCGTCTCAAGTTATGA CATGAACCGCGTCTCAAGTT ATGGCAACCGAGGAACCCATCATCG ATTGCGAAATTCATTGAAGAGAAA GCTTTGTCTTCGGCTTAGCAACTGTG CCTTTCTTCACCCCGGTAGCCTT TCGACCAAACAATTTTCGAGATG GAACACAAAACATCTTTACAATGCT GAACACAAAACATCTTTACAATGCTAAGG GTTATGCATGAACGTAATGCTC GCATGGTGGATTCACAATCCACT

Upstream non-coding region, − 243, forward Upstream non-coding region, − 149, forward Upstream non-coding region, − 373, forward Upstream non-coding region, − 373, forward Part of the region coding for N-terminal domain, 1, forward Part of the region coding for H1–5 globular domain, 349, forward Part of the region coding for C-terminal domain, 520, reverse Part of the region coding for C-terminal domain, 878, reverse Downstream non-coding region, 1620, reverse Downstream non-coding region, 932, reverse Downstream non-coding region, 932, reverse PsbA, forward trnH, reverse

PCR-amplification was carried out using BIS (produced by BIS-N, Koltsovo) and BIO-RAD MyCycler thermal cyclers under the following conditions: denaturation at 94 °C for 2 min 30 s followed by 5 cycles including denaturation at 94 °C for 30 s, annealing at 58 °C for 30 s, elongation at 70 °C for 1 min 30 s, followed by 35 cycles, including denaturation at 93 °C for 20 s, annealing at 56 °C for 30 s, elongation at 72 °C for 1 min 30 s. PCR reagents (Smart-Taq polymerase, dNTPs, buffers) were manufactured by Laboratory Medigen, Novosibirsk, the Taq polymerase used was also manufactured in Laboratory of Immunogenetics of ICG SD RAS, Novosibirsk. PCR-products were purified with Invisorb® Spin Filter PCRapid Kit according to manufacturer’s instructions. Purified PCRproducts were sequenced using Big Dye Terminators version 3.0 or 1.1 ABI PRISM at the sequencing centre ICG and IChBFM SD RAS. The sequence data were analysed using Staden package (Staden et al., 2000). 2.3. Data analysis Alignment of homologous nucleotide sequences was performed automatically with MEGA 4.0.2 package (Tamura et al., 2007) and MultAlin software (Corpet, 1988). Alignments were manually optimized. Diversity data analysis was carried out with MEGA 4.0.2 and DnaSP v5 packages (Librado and Rozas, 2009). Phylogenetic analysis was carried out for coding sequences of His5 (intron not included). All trees were rooted with the nucleotide sequences of paralogous pea histone H1 genes: His3 coding for H1 subtype 3 sequenced from accessions WL1238, VIR3971 (EMBL IDs: FR856880, FR856881 respectively) and an H1 gene sequenced from WL1446 (EMBL ID: FR856882) coding for an unidentified subtype allelic to PsH1b gene sequenced by Gantt and Key (1987) (EMBL ID: X05636). His5 and His3 are tightly linked at a distance ca 0.3 cM and reside in a tight, ca 1.5 cM long, group His(2–6) of 5 or 6 genes coding for H1 subtypes 2–6 (Trusov et al., 1994). Four types of phylogenetic reconstructions were derived using MEGA 4.0.2. package from His5 alignment for 1) all nucleotide substitutions, 2) non-synonymous nucleotide substitutions and 3) amino acid substitutions with Neighbour-Joining method, and 4) all nucleotide substitutions with Maximum Parsimony method. Evolutionary distances were computed by modified Nei–Gojobory method with Jukes– Cantor model for trees based on nucleotide substitutions. Amino acid sequence divergence values were calculated by Poisson correction model. For each phylogenetic reconstruction, 500 bootstrap replicates were made. Besides, the Maximum Likelihood bootstrap consensus tree was obtained with PHYLIP package (Felsenstein, 1989), also with 500 replicates. Phylogenetic reconstructions derived from PsbA-trnH spacer were generated by Neighbour-Joining method with 500 bootstrap replicates using MEGA 4.0.2. package.

3. Results 3.1. Sequencing His5 gene DNA sequence of His5 gene was identified in 65 pea lines (see Section 2.1 Plant Material). On the whole, 91 nucleotide substitutions in 87 polymorphic nucleotide sites and 36 alleles were found in the coding DNA sequences obtained (EMBL ID: FR846218, FR849610–FR849613, FR848833–FR848893). The mean nucleotide diversity Pi value was 0.0147, the haplotype diversity Hd was 0.956 (SD 0.016). Overwhelming majority of the derived amino acid sequences (58 of 65) are 256 amino acid residues long (Fig. s2). Deletions and insertions were found only in the C-terminal domain of H1–5. Identical insertion of 7 amino acid residues was found in accessions 703 and L95 (both P. fulvum). This insertion created a third copy of the KPKAAAA tandem repeat, starting after amino acid residue position 178 (hereinafter amino acid positions are given for the most common 256-amino acid length sequence). A deletion corresponding to 6 amino acid residues starting after the same position 178 was revealed in the accessions VIR3397, VIR6071, WL2140 and 702 (all P. fulvum). A different deletion of 5 amino acid residues starting at position 235 was observed in the accession VIR6560 (P. sativum subsp. sativum) originating from Tajikistan. In His-5 sequences without indels, the lengths of N-terminal, globular and C-terminal domains in amino acid residues were 50, 68, 129–142, respectively. On the whole, 45 variable amino acid positions were found in the reconstructed amino acid sequences of His5 products. 10 of them were localized in the N-terminal domain, 5 in the globular domain and 29 in the C-terminal domain. Using DnaSP v5 package for the entire set of His5 sequences obtained, the frequency of synonymous substitutions per synonymous site, Pi(s), was calculated, using Jukes and Cantor model, as 0.027; and that of non-synonymous substitutions per non-synonymous site, Pi(a), as 0.010. Application of MEGA 4.0.2 software to our sequences to calculate Z-test of selection at 500 bootstrap replicates (not shown) revealed non-neutral evolution regime almost exclusively for sequence pairs P. fulvum/other pea. DNA sequence coding for histone H1-5 were identical in 14 groups of more than one accession enumerated in Table 2. Of them, group 1 was of a special interest. It included full sequences from 6 accessions of remote provenance. Besides, partial sequences, obtained for accessions VIR3971 (Russia, Kirov Province), missing first 24 nucleotides after the start codon, VIR3913 (Tajikistan, Pamir), missing 39 nucleotides before the stop codon and VIR6135 (Greece), missing 32 nucleotides before the stop codon, were identical to them as well (unpublished, EMBL ID: FR849611–FR849613). This allele encodes a protein with a lysine residue substituted to asparagine and hence with a slowed mobility in the high resolution acetic acid/urea electrophoresis (see below). Group 4 included 7 accessions of wild and cultivated peas from remote regions, which was difficult to explain. It

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Table 2 Groups of pea accessions having identical nucleotide sequences of gene His5. No

Number of accessions

Taxonomical attribution, wild or cultivated

Accessions and their provenance

Marker combinations (A, B, C, D, rare) according to Kosterin and Bogdanova (2008)

1

6

P. sativum subsp. elatius (wild) P. sativum subsp. sativum (cultivated)

2

4

P. abyssinicum (cultivated)

A: VIR2524; B: Po14, P016, VIR4014 rare: VIR320* unidentified: VIR2514 A

3 4

2 5

5

3

P. sativum subsp. elatius (wild) P. sativum subsp. elatius (wild); P. sativum subsp. jomardii sensu Kosterin and Bogdanova (2008) (cultivated); P. sativum subsp. sativum (cultivated) P. sativum subsp. elatius (wild)

P016, Turkey, Denizli Province; VIR4014, Azerbaijan, Lenkoran; VIR2524, Israel, north Galilea; VIR320*, a genotype admixed to accession VIR320, ‘Palestine’. Pa014, Turkey, Tokat Province; VIR2514, Syria JI1876, Ethiopia, Makale; VIR2759, Ethiopia; VIR3567, Yemen, Dammar; WL1446, unknown. L100 (= 712), Israel, Be'er Sheva; 711, Israel, Jerusalem. 723, Sardinia; VIR2998, Georgia, WL805, Turkey VIR3424, VIR3439 — Egypt VIR4362, Russia, Vologda Province

6

2

P. sativum subsp. elatius (wild)

7

2

P. sativum subsp. elatius (wild)

8

2

P. sativum subsp. elatius (wild)

9

3

P. sativum subsp. elatius (wild)

10 11

3 2

12

2

13

2

14

4

P. sativum P. sativum (wild) P. sativum (wild) P. sativum (wild) P. sativum P. fulvum

subsp. elatius (wild) subsp. elatius subsp. elatius subsp. elatius subsp. transcaucasicum (cultivated)

JI1094, JI1095 — Greece, Mt AthosPeninsula; JI3553, France, Marseille environs WL2123, Israel, Jordan valley; 722 Israel, Adamit VIR320, ‘Palestine’, 714, Israel, between Bet Shemesh and Bet Gurvin PI343993, Turkey, 5 km SW Kale; VIR7327, Turkey JI1092, JI2055 — Greece, Mt. Athos peninsula; PI344537, Italy, Sicily, Campania, Alburni Mt. JI1091, JI1096, PI344008 — Greece, Mt. Athos peninsula JI1794, Israel/Syria, Holan Heights; P012 Turkey, Adiyamen Province CE1, Ukraine, Crimea, Simeiz; CE2, Ukraine, Crimea, Kara Dag VIR1851, Georgia VIR3249, Georgia VIR2523, ‘Palestine’; VIR6071, Israel, 30 km SW Jerusalem; JI2140, Israel, Valley of the Cross; 702, Israel, Mt. Canaan

cannot be excluded that some contamination occurred during cultivation in germplasm collections. 3.2. Phylogenetic analysis of His5 sequences We obtained three phylogenetic reconstructions for His5 coding sequence basing on the distance, parsimony and maximum likelihood analyses (Figs. 1–2, s3). The outgroup used, being two paralogous H1 genes, appeared somewhat too distant, so that the branches leading to it and the His5 tree were too long for an in scale illustration. So, in Fig. 1 the outgroup is not shown (the length of the branch leading to the His5 subtree is provided in the legend), while Figs. 2 and s3 show only tree topology. We also obtained Neighbour-Joining phylogenetic trees basing separately on synonymous, non-synonymous and amino acid substitutions (not shown). They were less informative, especially that for synonymous substitutions; as only the P. fulvum cluster had on the latter a support greater than on the analysed tree for all nucleotide substitutions (78 vs 53). On the tree constructed by the Neighbour-Joining method (further in the text ‘NJ tree’) (Fig. 1), the first dichotomy separated P. fulvum from other accessions, although bootstrap value was not very high (53 and 57 for P. fulvum cluster and for the rest of accessions respectively). The cluster that hosts P. sativum appeared rather unstructured with many branches formed by carriers of combinations A and C. Most representatives of combination C originating from Central Mediterranean (Greece, France and Italy) reside in a weakly supported (bootstrap value 32) branch, which also included a well supported cluster of all accessions of P. abyssinicum. Long branches with significant bootstrap values (more than 90) united different accessions from Israel mostly with combination A. Combination A appeared to be ancestral for the genus and C is its earliest derivative.

A A: 723; B: VIR2998, VIR4362; rare: WL805; D: VIR3424, VIR3439 C A: 722; rare: WL2123 A C: PI343993; D: VIR7327 C: JI1092, JI2055; D: PI344537 C B B B

A

Among those branches, however, there was a large and well supported (bootstrap value 96) branch which harboured wild and cultivated P. sativum accessions representing mainly evolutionary younger combinations D and B, further in the text called the ‘D–B clade’. The only accession carrying one of the younger combinations, namely D, found outside this clade, was PI344537, which appeared to relate closer to accessions with combination C from Central Mediterranean. However, 3 accessions carrying combination A were found inside the D–B clade: VIR2521, VIR2524 from Israel and 723 from Sardinia, as well as accession PI343993 with combination C from Turkey. The D–B clade hosted most representatives of ‘rare’ combinations included in this study (3 out of 4: VIR320* from Palestine, WL805 and Ps008 from Turkey). Most of the wild accessions found in this clade represented either Turkey or Tauro-Caucasian area, although 4 accessions from Israel were also found there. Inside the D–B clade, there was a small cluster that harboured accessions Ps002, P012 and JI1794, all three being wild P. sativum subsp. elatius with combination B, the two latter with identical His5 sequences. Noteworthy, JI1794 possesses the same karyotype as the cultivated pea (Ben-Ze'ev and Zohary, 1973). The topology of the tree constructed by the maximum parsimony method (further ‘MP tree’) (Fig. s3) resembled greatly that of NJ tree and even bootstrap values differed insignificantly. The main differences were the position of accession P013 (wild P. sativum from Turkey) and the bootstrap values of the Ps002, P012, JI1794 and VIR7327, PI343993, VIR6560 clusters. Accession P013 appeared as a separate branch in the very base of the P. sativum cluster on NJ tree with the bootstrap support of 57 while on MP tree it clustered with accessions 713, 714 and VIR320, although the bootstrap values were lower than 20. The bootstrap values for the VIR7327, PI343993, VIR6560 cluster were 60 on NJ tree while on MP tree these accessions were located inside a larger set of accessions, which included JI2105 and P017. However the bootstrap support for this cluster was weak. The cluster that hosts accessions Ps002,

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Fig. 1. Neighbour-joining tree of the studied accessions of genus Pisum L. based on His5 sequences. The outgroup is not shown as it was too long in comparison with the His5 subtree. The outgroup included paralogous pea H1 genes: sequences from WL1238 and VIR3971 of the gene coding for H1–3 subtype and a sequence from WL1446 of the PsH1b gene. The distance between the outgroup bifurcation and the root of the His5 subtree is estimated to be 0.057. Taxonomical attribution and combinations (A, B, C, D, rare, see Kosterin et al., 2010) of the three markers RbcL, Cox1 and SCA are indicated. Provenance of accessions of P. sativum subsp. elatius is indicated.

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Fig. 2. Maximum likelihood bootstrap consensus tree of the studied accessions of genus Pisum L. based on His5 sequences. Designations as in Fig. 1.

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P012 and JI1794, which was weakly supported on NJ tree, existed on MP tree with bootstrap of 71. The Maximum Likelihood bootstrap consensus tree (further ML tree) (Fig. 2) was consistent with the two other reconstructions: tree topology and bootstrap values were basically the same. Notably, ML tree demonstrated the highest bootstrap value for P. fulvum cluster, 83. The position of the accession VIR6560 was similar to that on MP tree rather than on NJ tree. The position of P013 was uncertain but clearly it was not in the base of P. sativum subtree. The bootstrap support for the Ps002, P012 and JI1794 cluster was 84. 3.3. Comparison of His5 and PsbA-trnH utility for phylogenetic analysis To evaluate the utility of the His5 gene for phylogenetic inference we sequenced the PsbA-trnH spacer (EMBL ID: FR856864–FR856878, HE601757–HE601760; Fig. s4) in a set of 19 accessions including 4 P. fulvum accessions (VIR6070, L95, 707, 708), 2 P. abyssinicum (JI1876, WL1446), 9 wild P. sativum subsp. elatius VIR320, VIR2998, JI1092, JI1794, PI344537, CE1, P015, P017, P008, and 4 cultivated P. sativum subsp. sativum (WL1238, VIR3249, VIR3439, VIR6560). We found only 2 polymorphic sites, one of which corresponded to a nucleotide substitution C → T in the position 64 from the beginning of the spacer found in both studied P. abyssinicum sequences, the second corresponded to a nucleotide substitution A → T in the position 128 shared by accessions P017 and JI7194. A 8 bp deletion embracing nucleotides 142–149 was revealed in the cultivated accessions VIR6560, WL1238 (P. sativum subsp. sativum), VIR3439 (P. sativum subsp. jomardii), VIR3249 (P. sativum subsp. transcaucasicum) as well as in 1 ambiguously wild accession VIR2998 (? P. sativum subsp. elatius). We compared nucleotide sequence diversity parameters of PsbAtrnH to that of His5 in the same set of 19 accessions (Table 3) and obtained NJ trees basing on these two markers (supplementary Fig. s5). The His5 tree for this set of accessions was well-structured and resolved all main clusters that appeared in the trees for larger set of accessions based on His5 gene, while variation of PsbA-trnH was insufficient to reconstruct phylogeny. 4. Discussion 4.1. His5 gene variation Histones (except for H4) are roughly classified into replication dependent, replication independent or replacement, and tissue specific; with the replication dependent histone genes arranged into repeat clusters and lacking introns and the replication independent histone genes found outside clusters and containing introns (Rooney et al., 2002). However in plants, clusters of replication dependent histone genes may be small, if any, while H1 genes are not clustered (Eirín-López et al., 2004). Noteworthy, all the seven subtypes of histone H1 in pea are encoded by unique genes (Kosterin et al., 1994), so far sequenced H1 genes contain intron, so they show features of replication independent histone genes, although subtype H1–7 is present only in actively growing tissues (Kosterin et al., 1994). In this paper, we analysed variation among alleles of one of these paralogous genes, His5, which is a member of a loose (ca 1.5 cM) group of 5 linked quite divergent H1 genes, His(2–6), which are not repeats (Trusov et al., 1994). Intergene conversion or recombination

Table 3 Diversity parameters of the PsbA-trnH spacer and His5 sequences.

Number of sequences Number of sites Number of polymorphic sites Average number of differences Nucleotide diversity, Pi

involving these paralogs cannot be excluded but must be extremely rare; so far we faced no sign of it. Besides, intragene recombination has a little effect on sequence variation of His5 alleles in pea which is predominantly a self-pollinator hence highly homozygous. Earlier (Berdnikov et al., 1993a; Kosterin et al., 1994) variation in pea histone H1 was investigated using high resolution acetic acid/urea PAGE (Panyim and Chalkley, 1969). In these denaturing conditions, protein mobility depends strongly and positively on the number of positively charged amino acid residues and mildly and negatively on the total number of amino acid residues and can be calculated by an empirical formula (Berdnikov et al., 1993b), verified for the here studied H1 subtype 5 electromorphs in Bogdanova et al. (2005). In cultivated peas, a limited number of distinct electromorphs was observed (Berdnikov et al., 1993a), while in wild peas there were more electromorphs, sometimes differing too subtly to estimate their number (Kosterin et al., 1994). Sequencing of H1 genes showed that equal electrophoretic mobility can be achieved by different amino acid substitutions and/or indels. For instance, the same gain in electrophoretic mobility would result from different deletions in the C-terminal domain, KPKAVA in 4 accessions of P. fulvum and SVKAK in accession VIR6560 of P. sativum subsp. sativum from Tajikistan, as well as from the substitution of glycine to lysine in the globular domain in accession P017 of P. sativum subsp. elatius from Turkey. The same loss in electrophoretic mobility would be due to the insertion KPKAAAA in accessions 703 and L95 of P. fulvum and a substitution of lysine to asparagine in 10 accessions of P. sativum. This group of 10 accessions, originating from different parts of the world (Israel, Syria, Turkey, Azerbaijan, Greece, Tajikistan, North Russia), both wild and cultivated, were found to have the identical His5 sequence (6 complete sequences, 4 partial). A broad distribution of an identical allele suggests its rapid spread. This allele encodes the electrophoretic “variant 1” the frequencies of which among aboriginal landraces of cultivated peas were found to be the greatest in North Russia and highlands of Central Asia and showed a strong negative correlation with the accumulated temperatures of the vegetation period (Berdnikov et al., 1993a). This may be interpreted so that natural selection, acting during primitive cultivation in habitats with cold climate, aided in expansion of this allele over remote regions (Berdnikov et al., 1993a). All four P. abyssinicum accessions involved carried identical His5 sequences. This agrees with a very recent radiation of this taxon estimated as recent as 4000 years (Ellis et al., 1998). In this case, variation in His5 was nil and hence insufficient for estimations. The data on synonymous and non-synonymous substitutions suggests some non-neutral evolution, quite expectable in a functional protein. Z-test revealed non-neutral regime in cases when sequences of P. fulvum were compared to those of other peas, that is only P. fulvum is divergent enough to make it noticeable. Indels were found only in the region coding for C-terminal domain of H1-molecules. Among 12 accessions of P. fulvum we found an insertion (2 accessions) and a deletion (4 accessions), versus 1 deletion among 53 accessions of P. sativum subsp. sativum. This asymmetry may be explained if we assume that P. fulvum existed and accumulated variation at the place of its origin while P. sativum still in its wild state underwent at least two expansions, from east to west and then an almost backward from west to north-east (Kosterin et al., 2010). During this migration it might have lost some of its variation because of the bottleneck effects. 4.2. Phylogenetic trees

PsbA-trnH

His5

19 201 2 0.40 0.00198

19 792 52 11.25 0.01459

Pisum L. is a small genus, represented by three very close species, that belongs to the tribe Fabeae (syn. Vicieae). On phylogenetic reconstructions of this tribe based on several chloroplast genes and nuclear internal transcribed spacer, Pisum appears monophyletic (holophyletic) and a sister group to the monotypical genus Vavilovia Fed., together with which it forms a branch residing inside the genus Lathyrus L.,

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hence paraphyletic, which, in turn, resides inside the also paraphyletic genus Vicia L. (Kenicer et al., 2005, 2009a, b; Smýkal et al., 2010). All phylogenetic studies, including ours, agree in that the morphologically peculiar P. fulvum forms the most distinct basal clade (Hoey et al., 1996; Jing et al., 2007; Vershinin et al., 2003). At the same time P. abyssinicum, morphologically very close to P. sativum, forms a separate branch among wild P. sativum accessions. So, acceptance of P. abyssinicum makes the remaining species P. sativum paraphyletic, therefore it does not fit a strictly phylogenetic taxonomical system. However it fits the biological species concept in a sense of E. Mayr (1942), based on reproductive integrity and reproduction isolation, since abyssinicum is well isolated from other peas due to chromosome rearrangements (Conicella and Errico, 1990) and additional severe sterility of hybrids due to some conflict of the parental genetic programmes (unpublished). The biological species concept can be safely applied to peas, which are not perfect self-pollinators and gene exchange is substantial from the evolutionary point of view (Bogdanova and Berdnikov, 2000; Jing et al., 2007; Loenning, 1984). On the reconstructions presented in (Vershinin et al., 2003), P. abyssinicum formed a clade distant from that of other cultivated peas (P. sativum subsp. sativum s. l.), suggesting that it was domesticated independently. Our study supports this position as P. abyssinicum appears closer to some wild P. sativum forms and shares with them the plesiomorphic, ancestral for the genus marker combination A, while P. sativum subsp. sativum s. l. possesses the apomorphic combination B. Vershinin et al. (2003) indicated that ‘P. abyssinicum displays extreme homogeneity possibly due to a bottleneck effect’. Our study shows that all 4 investigated P. abyssinicum accessions possess identical His5 sequences, in accordance with the latter assumption. So, the two cultivated lineages, P. abyssinicum and P. sativum subsp. sativum, were derived from different wild forms of P. sativum L. Early divergence within P. sativum is poorly resolved in all phylogenetic trees, including ours, which contain many subtle but ancient lines. In this situation, for practical reason it seems worthy to avoid isolation of these lines into separate taxa and to artificially ascribe all them to the paraphyletic P. sativum subsp. elatius (Bieb.) Schmahl, as presently widely accepted. In our phylogenetic trees obtained by different methods, among basic branches, represented by accessions with combinations A and C, that are either poorly resolved or small, a robust and well supported cluster is found which is formed mostly by accessions with combinations B and D, ‘D–B clade’. Most of the ancient branches in the base of the trees lead to accessions from Israel with combination A. Relationships between accessions with combinations A and C from Central and West Mediterranean, namely from Greece, Italy, France and Spain, are resolved more poorly. Probably, this is due to their recent radiation in contrast to older lineages from Eastern Mediterranean. Inside the ‘D–B clade’, a separate branch is formed by three wild accessions: JI1794 (Holan Heights), Ps002 and P012 (Turkey). BenZe'ev and Zohary (1973) attributed JI1794 to ‘northern humile’, that is, wild pea with the karyotype typical for the cultivated pea. Other wild accessions in the ‘DB-clade’ cluster originate mostly from Turkey or Tauro-Caucasian area. Nevertheless, 4 wild accessions from other regions with ancestral combinations A and C are also found in ‘B–D cluster’. Two of them, VIR2524 and VIR 2521, both P. sativum subsp. elatius with combination A from Israel, were received from AllUnion Institute of Plant Breeding (VIR) as highly heterogeneous that most probably resulted from contamination while reproducing the collection. Noteworthy, VIR2521 carries His5 allele the frequency of which in regional samples of cultivated peas correlates with the accumulated temperatures of the vegetation period (Berdnikov et al., 1993a). This variant is supposed to have spread over cultivated peas quite recently. Accession 723 from Sardinia with combination A resembles cultivated peas in its habitus, that may result from some introgression. Accession PI343993 carrying combination C comes from Turkey where representatives of combinations A and B coexist, so it is likely a hybrid form.

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Jing et al. (2007) undertook a combined phylogenetic analysis of 39 pea genes altogether and demonstrated that a single gene analysis can lead to a distorted view. Therefore, our results should be interpreted with caution. Nevertheless, the phylogenetic trees based on His5 sequences quite adequately reflect the current pea taxonomy: we observe distinct and clear-cut clade of P. fulvum, also a distinct clade of P. abyssinicum inside heterogeneous wild representatives of P. sativum. Of the published molecular phylogenetic studies, the set of accessions used in (Ben-Ze'ev and Zohary, 1973) was involved only into the work by Hoey et al. (1996), in which the only clear-cut cluster observed on all reconstructions was formed by ‘southern humile’, while the relationships between ‘elatius’, ‘northern humile’ and P. sativum were controversial. Other studies (Ellis et al., 1998; Jing et al., 2007; Vershinin et al., 2003) involved much greater sets of accessions but unfortunately not that set. In these studies, the phylogenetic relationships between diverse forms of P. sativum remained mostly unresolved and could not be coordinated with karyological classes by (Ben-Ze'ev and Zohary, 1973). Our data, together with the results on the three markers from different cellular genomes (Kosterin and Bogdanova, 2008; Kosterin et al., 2010), resolve an important evolutionary event within P. sativum (equivalent to the ‘humile–elatius–sativum complex’), that is, separation of a robust monophyletic D–B clade which hosts both wild and cultivated peas bearing marker combinations D and B. Our study involved the accessions by (Ben-Ze'ev and Zohary, 1973) that allowed us to associate the D–B clade with one of the two karyological classes isolated by the last cited authors, namely that including cultivated peas and ‘northern humile’. This clade resides among many other, more subtle branches representing wild forms of P. sativum, mostly with combinations A or C, including the cluster of P. abyssinium, which may be called ‘basal A– C group’. The origin of the robust ‘D–B clade’ should have more or less coincided with the transition C→D, that is, loss of the recognition site for the restriction endonuclease AspLEI in the plastid gene RbcL due to a synonymous substitution T→C (Kosterin and Bogdanova, 2008). This transition should have taken place somewhere in South Europe, since representatives of combination C are most frequent in the Iberian, Apennine and Balkan peninsulas while their remote descendants with combination B occupy the Tauro-Caucasian region (Kosterin et al., 2010). A more precise location is impossible since 6 known carriers of combination D are scattered over Mediterranean. As well they are scattered without a clear pattern among branches of the ‘D–B clade’, so we cannot resolve the D→B transition, due to mutation in SCA, either on the phylogenetic trees or on the geographical map. It is not excluded that all D-carriers resulted from secondary hybridisation of peas with combinations A or C with those with combination B, which could take place even during reproduction of germplasm collections. The A→C transition cannot be indicated on our phylogenetic trees since the topology of branches with combinations A and C is not well resolved. Noteworthy, the NJ tree contains a weakly supported (bootstrap value 32) branch comprising most representatives of combination C originating from Central and West Mediterranean and all accessions of P. abyssinicum (Fig. 1). Hence, within the ‘basal A–C group’, this taxon may be more related to the combination C-carriers which we suppose to have given rise to the ‘D–B clade’. There is no doubt that the cultivated P. sativum subsp. sativum arose by domestication within the ‘D–B clade’. However, its representatives do not form a clade of their own and reside side-by-side with wild forms. This may be explained by either insufficient phylogenetic signal from variation within His5 or by secondary gene exchange within wild and cultivated representatives of the ‘D–B clade’. The latter is supported by the occurrence of two representatives of combination A (723 and VIR2524, the latter proved to be heterogeneous) and one of combination C (PI344993) within the ‘D–B clade’ and one representative of combination D (PI344537) within the ‘basal A–C group’.

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4.3. Variation of the PsbA-trnH plastid intergenic spacer in pea For comparison we chose the plastid intergenic spacer PsbA-trnH as a marker popular in interspecies studies, which is rather variable and was proposed as a candidate for molecular barcoding in plants, which faces more difficulties than in animals in finding a marker that both can differentiate species and is easily sequenced from the same set of primers in the majority of taxa (Kress and Erickson, 2007; Kress et al., 2005; Sass et al., 2007). Recent studies indicate that PsbA-trnH is one of the most variable regions in plant genome and is easily amplified. Its length on average is about 450 bp (Kress et al., 2005). In a comparative study, PsbA-trnH was successfully amplified for all eight investigated genera and 19 species while other candidate markers, including ITS, could not be amplified in one or more taxa (Kress et al., 2005). PsbA-trnH had the highest divergence value in six of the eight genera and in 11 of the 14 species pairs, compared with other plastid regions investigated. The length of the intergenic spacer PsbA-trnH ranged from 119 to 1094 bp across 53 families of flowering plants and all investigated species had unique PsbA-trnH spacer sequences (Kress et al., 2005). It turned out that the PsbA-trnH spacer in pea is only 201 bp long. We found only 2 polymorphic sites and a 8 bp deletion in VIR2998, VIR6560, VIR3439, VIR3249 and WL1238. This deletion seems to be a marker of the cultivated P. sativum subsp. sativum and P. sativum subsp. jomardii. A wild carrier of combination D, PI344537 does not have this deletion. So far the only wild accession carrying it is VIR2998 of dubious origin: its original provenance was stated to be Georgia, but the samples were obtained by VIR from the Riga botanical garden in Latvia. This accession was also heterogeneous that could be explained by contamination through cross-pollination during cultivation in germplasm collection. As this deletion is not shared by wild peas and is found in both P. sativum subsp. sativum and P. sativum subsp. jomardii, it is logical to conclude that it arose around the time of their domestication and that P. sativum subsp. sativum and P. sativum subsp. jomardii are unlikely to have been domesticated independently. P. sativum subsp. jomardii accessions possess combination D while all tested P. sativum subsp. sativum, but 2 accessions carry combination B (Kosterin and Bogdanova, 2008; Kosterin et al., 2010). Earlier on this basis we suggested that P. sativum subsp. jomardii originated from some wild pea population carrying combination D, while P. sativum subsp. sativum descend from some other wild population carrying combination B (Kosterin and Bogdanova, 2008). However, the common deletion in the PsbA-trnH spacer disproves this supposition. ‘P. sativum subsp. jomardii’ seems to be just a local cultivated form of P. sativum subsp. sativum confined to Egypt. Most probably, P. sativum subsp. jomardii acquired combination D from cross-pollination of P. sativum subsp. sativum with wild carriers of either combination A or combination C. From this point of view, ‘P. sativum subsp. jomardii’ does not deserve a subspecies status proposed in (Kosterin and Bogdanova, 2008). Apparently, PsbA-trnH in pea is not variable enough to resolve phylogeny of the genus. It allows to distinguish cultivated forms of P. sativum subsp. sativum plus ‘P. sativum subsp. jomardii’ with the 8 bp deletion as well as the independently domesticated P. abyssinicum having a nucleotide substitution. However, there is no difference in the PsbA-trnH spacer sequence between wild P. sativum and P. fulvum, which are clearly distinct species. Contrary, a histone H1 gene, at least His5 and at least in pea, allowed to reconstruct a phylogenetic tree even from a small set of accessions. This is not because PsbA-trnH is too short in pea, for we see that the number of polymorphic sites per length unit is 6 times lower in PsbA-trnH than in His5. Indeed, the PsbA-trnH spacer appears to scarcely fit phylogenetic studies in legumes and some other markers from the same plastid genome also suggested for barcoding in plants provide better results in this family, namely RbcL (Doyle et al., 1997; Kass and Wink, 1996; Lavin et al., 2005), MatK (Lavin et al., 2005; Steele and Wojciechowski, 2003; Wojciechowski et al., 2004), trnL-F and trnS-G

(Kenicer et al., 2005), the three latter being applied to the genus Lathyrus which is the closest relative of Pisum. 4.4. Concluding remark His5 encodes rather a minor subtype, H1–5, and may be less apt to purifying selection and hence less conservative than paralogous genes for some other, more abundant in pea H1 subtypes, such as H1–1 and H1–2, or for a more specialized subtype H1–7, present only in actively growing tissues (Bogdanova et al., 2007; Kosterin et al., 1994). To sequence other paralogous H1 in pea and to compare results with those reported here is a matter of further research. We have shown that the gene of an H1 subtype offered useful phylogenetic information at the intrageneric and even intraspecies level. However, usability of this very gene and our experimental design is limited to peas since the primers we use are too specific to the noncoding regions adjacent to His5. Even other legumes of the tribe Fabaeae may not have the ortholog of this very gene, although they obviously have a variable number of H1 subtypes orthologous to subtypes 1, 6, 7 and the group of subtypes 2–5 in pea (Kosterin et al., 1994). By now in pea, we sequenced the genes for subtype 1 (Berdnikov et al., 2003), subtype 3 (used as an outgroup in this paper), subtype 4 (unpublished), subtype 5 (Bogdanova et al., 2005 and this paper), subtype 6 (in preparation) and subtype 7 (Bogdanova et al., 2007). Their sequences are quite divergent so that they can be easily distinguished, while their non-coding neighbourhoods allow specific primers. Only in case of the subtype 4 gene we faced some obvious cases of cross-amplification with the subtype 3 gene (unpublished). Using specific primers to the non-coding regions of the gene for pea subtype 6, we succeeded in specific amplification of its ortholog in Vicia unijuga A. Br. (in prep.) but not in cases of some other H1 genes and legume species. Hence, at present histone H1 genes cannot be easily used as convenient phylogenetic markers at above-species level due to the lack of universal primers. However, some experimental approaches basing on the conserved histone H1 globular domain can be worked out which would allow a wider usability of histone H1 genes as phylogenetic markers. Acknowledgment This work was supported by the Russian Foundation for Fundamental Research, Grant 10-04-00230-а. We are grateful to the three anonymous Referees whose valuable comments allowed to greatly improve the paper. Appendix A. Supplementary data Supplementary data to this article can be found online at http:// dx.doi.org/10.1016/j.gene.2012.05.026. References Alami, R., et al., 2003. Mammalian linker-histone subtypes differentially affect gene expression in vivo. Proc. Natl. Acad. Sci. 100, 5920–5925. Allan, J., Mitchell, N., Harborne, C., Bohm, C., Crane-Robinson, C., 1986. Roles of H1 domains in determining higher order chromatin structure and H1 location. J. Mol. Biol. 187, 591–601. Ben-Ze'ev, N., Zohary, D., 1973. Species relationship in the genus Pisum L. Isr. J. Bot. 22, 73–91. Berdnikov, V.A., Bogdanova, V.S., Rozov, S.M., Kosterin, O.E., 1993a. The geographic patterns of histone H1 allelic frequencies formed in the course of pea (Pisum sativum L.) cultivation. Heredity 71, 199–209. Berdnikov, V.A., Rozov, S.M., Temnykh, S.V., Gorel', F.L., Kosterin, O.E., 1993b. Adaptive nature of interspecies variation of Histone H1 in insects. J. Mol. Evol. 36, 497–507. Berdnikov, V.A., Gorel, F.L., Bogdanova, V.S., Kosterin, O.E., Trusov, Y.A., Rozov, S.M., 1999. Effect of a substitution of a short chromosome segment carrying a histone H1 locus on expression of the homeiotic gene Tl in heterozygote in the garden pea Pisum sativum L. Genet. Res. 73, 93–109. Berdnikov, V.A., Gorel, F.L., Bogdanova, V.S., Kosterin, O.E., Trusov, A.Y., Rozov, S.M., 2003. Large changes in the structure of the major histone H1 subtype result in small effects on quantitative traits in legumes. Genetica 119, 167–182.

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