Orcein

The taxa Polyommatus (Agrodiaetus) shirkuhensis (Iran, Yazd Province, Shirkuh Mts., Deh-Bala village, 2900-3150 m, 12 July 2005, samples J299-1, J299-2 and J299-3, J302 and J304) and Polyommatus (Agrodiaetus) bograbirjandensis(Iran, South Khorasan Province, 26 km N of Birjand, 1900-2000 m, 14 July 2005, samples J305, J306, J307, J307-1, J307-2, J307-3, J307-4, J315, J318 and J319) were collected exactly in their type localities.
Fresh (not worn) adult males were used to investigate the karyotypes. After capturing a butterfly in the field, it was placed in a glassine envelope for 1-2 hours to keep it alive until we processed it. Testes were removed from the abdomen and placed into a small 0.5 ml vial with a freshly prepared fixative (ethanol and glacial acetic acid, 3:1). Then each wing was carefully removed from the body using forceps. The wingless body was placed into a plastic, 2 ml vial with pure 96% ethanol. The samples are kept in the Zoological Institute of the Russian Academy of Sciences.
Testes were stored in the fixative for 1-12 months at +4°C. Then the gonads were stained in 2% acetic orcein for 30-60 days at +18-20°C. Different stages of male meiosis were examined by using a light microscope Amplival, Carl Zeiss. We have used an original two-phase method of chromosome analysis (Lukhtanov and Dantchenko 2002 (link), Lukhtanov et al. 2006 ).
A 643 bp fragment of mitochondrial gene PageBreakcytochrome oxidase subunit I (COI) and 592 bp fragment of nuclear internal transcribed spacer 2 (ITS2) were used to analyze clustering of the specimens. Primers and the protocol of DNA amplification were given in our previous publication (Lukhtanov et al. 2008 ). The sequences were edited and aligned using BioEdit 7.0.3 (Hall 1999 ). Since Polyommatusicarus (Rottemburg, 1775) and Polyommatusstempfferi (Brandt, 1938) were earlier inferred as outgroups to the subgenus Agrodiaetus (Talavera et al. 2013 ), we used them to root the phylograms.
Sequences of the following additional representatives of the subgenus PageBreakAgrodiaetus were found in GenBank (Wiemers 2003 , Wiemers and Fiedler 2007 , Wiemers et al. 2009 (link), Kandul et al. 2004 (link), 2007 (link), Lukhtanov et al. 2005 (link)) and used for phylogenetic inference: Polyommatus (Agrodiaetus) ainsae (Forster, 1961), Polyommatus (Agrodiaetus) achaemenes Skala, 2002, Polyommatus (Agrodiaetus) actinides (Staudinger, 1886), Polyommatus (Agrodiaetus) admetusmalievi (Dantchenko et Lukhtanov, 2005), Polyommatus (Agrodiaetus) aereus Eckweiler, 1998, Polyommatus (Agrodiaetus) alcestiskaracetinae (Lukhtanov et Dantchenko, 2002), Polyommatus (Agrodiaetus) altivagans (Forster, 1956), Polyommatus (Agrodiaetus) antidolus (Rebel, 1901), Polyommatus (Agrodiaetus) ardschira (Brandt, 1938), Polyommatus (Agrodiaetus) baltazardi (de Lesse, 1963), Polyommatus (Agrodiaetus) baytopi (de Lesse, 1959), Polyommatus (Agrodiaetus) bilgini (Dantchenko et Lukhtanov, 2002), Polyommatus (Agrodiaetus) birunii Eckweiler et ten Hagen, 1998, Polyommatus (Agrodiaetus) caeruleus (Staudinger, 1871), Polyommatus (Agrodiaetus) carmoncarmon (Herrich-Schäffer, 1851), Polyommatus (Agrodiaetus) carmonmunzuricus (Rose, 1978), Polyommatus (Agrodiaetus) ciscaucasicus (Forster, 1956), Polyommatus (Agrodiaetus) cyaneus (Staudinger, 1899), Polyommatus (Agrodiaetus) dagestanicus (Forster, 1960), Polyommatus (Agrodiaetus) dagmara (Grum-Grshimaïlo, 1888), Polyommatus (Agrodiaetus) damocles (Herrich-Schäffer, 1844), Polyommatus (Agrodiaetus) damon (Dennis et Schiffermüller, 1775), Polyommatus (Agrodiaetus) damonealtaicus (Elwes, 1899), Polyommatus (Agrodiaetus) damonedamone (Eversmann, 1841), Polyommatus (Agrodiaetus) damoneirinae (Dantchenko, 1997), Polyommatus (Agrodiaetus) dantchenkoi Lukhtanov et Wiemers, 2003, Polyommatus (Agrodiaetus) demavendi (Pfeiffer, 1938), Polyommatus (Agrodiaetus) dizinensis (Schurian, 1982), Polyommatus (Agrodiaetus) dolusvittata (Oberthür, 1892), Polyommatus (Agrodiaetus) ectabanensis (de Lesse, 1964), Polyommatus (Agrodiaetus) elbursicus (Forster, 1956), Polyommatus (Agrodiaetus) eriwanensis (Forster, 1960), Polyommatus (Agrodiaetus) erschoffii (Lederer, 1869), Polyommatus (Agrodiaetus) faramarzii Skala, 2001, Polyommatus (Agrodiaetus) femininoides (Eckweiler, 1987), Polyommatus (Agrodiaetus) firdussii (Forster, 1956), Polyommatus (Agrodiaetus) fulgens (Sagarra, 1925), Polyommatus (Agrodiaetus) glaucias (Lederer, 1870), Polyommatus (Agrodiaetus) gorbunovi (Dantchenko et Lukhtanov, 1994), Polyommatus (Agrodiaetus) haigi (Dantchenko et Lukhtanov, 2002), Polyommatus (Agrodiaetus) hamadanensis (Lesse, 1959), Polyommatus (Agrodiaetus) hopfferi (Gerhard, 1851), Polyommatus (Agrodiaetus) huberti (Carbonell, 1993), Polyommatus (Agrodiaetus) iphidamon (Staudinger, 1899), Polyommatus (Agrodiaetus) iphigenia (Herrich-Schäffer, 1847), Polyommatus (Agrodiaetus) iphigenides (Staudinger, 1886), Polyommatus (Agrodiaetus) karatavicus Lukhtanov, 1990, Polyommatus (Agrodiaetus) karindus (Riley, 1921), Polyommatus (Agrodiaetus) kendevani (Forster, 1956), Polyommatus (Agrodiaetus) kermansis (de Lesse, 1963), Polyommatus (Agrodiaetus) khorasanensis (Carbonell, 2001), Polyommatus (Agrodiaetus) klausschuriani ten Hagen, 1999, Polyommatus (Agrodiaetus) kurdistanicus (Forster, 1961), Polyommatus (Agrodiaetus) lorestanus Eckweiler, 1997, Polyommatus (Agrodiaetus) lukhtanovi (Dantchenko, 2005), Polyommatus (Agrodiaetus) luna Eckweiler, 2002, Polyommatus (Agrodiaetus) magnificus (Grum-Grshimaïlo, 1885), Polyommatus (Agrodiaetus) masulensis ten Hagen et Schurian, 2000, Polyommatus (Agrodiaetus) mediator (Dantchenko et Churkin, 2003), Polyommatus (Agrodiaetus) menalcas (Freyer, 1837), Polyommatus (Agrodiaetus) merhaba De Prins, van der Poorten, Borie, van Oorschot, Riemis et Coenen, 1991, Polyommatus (Agrodiaetus) mithridates (Staudinger, 1878), Polyommatus (Agrodiaetus) mofidii (de Lesse, 1963), Polyommatus (Agrodiaetus) ninae (Forster, 1956), Polyommatus (Agrodiaetus) peilei (Bethune-Baker, 1921), Polyommatus (Agrodiaetus) pfeifferi (Brandt, 1938), Polyommatus (Agrodiaetus) phyllides (Staudinger, 1886), Polyommatus (Agrodiaetus) phyllis (Christoph, 1877), Polyommatus (Agrodiaetus) pierceae (Lukhtanov et Dantchenko, 2002), Polyommatus (Agrodiaetus) poseidon (Herrich-Schäffer, 1851), Polyommatus (Agrodiaetus) poseidonides (Staudinger, 1886), Polyommatus (Agrodiaetus) pulcher (Sheljuzhko, 1935), Polyommatus (Agrodiaetus) putnami (Dantchenko et Lukhtanov, 2002), Polyommatus (Agrodiaetus) ripartii (Freyer, 1830), Polyommatus (Agrodiaetus) ripartiiparalcestis (Forster, 1960), Polyommatus (Agrodiaetus) rjabovi (Forster, 1960), Polyommatus (Agrodiaetus) rovshani (Dantchenko et Lukhtanov, 1994), Polyommatus (Agrodiaetus) sennanensis (de Lesse, 1959), Polyommatus (Agrodiaetus) shahkuhensis (Lukhtanov, Shapoval et Dantchenko, 2008), Polyommatus (Agrodiaetus) shahrami Skala, 2001, Polyommatus (Agrodiaetus) shamil (Dantchenko, 2000), Polyommatus (Agrodiaetus) sorkhensis Eckweiler, 2003, Polyommatus (Agrodiaetus) surakovi (Dantchenko et Lukhtanov, 1994), Polyommatus (Agrodiaetus) tankeri (de Lesse, 1960), Polyommatus (Agrodiaetus) tenhageni Schurian et Eckweiler, 1999, Polyommatus (Agrodiaetus) transcaspica (Heyne, 1895), Polyommatus (Agrodiaetus) turcicolus (Koçak, 1977), Polyommatus (Agrodiaetus) turcicus (Koçak, 1977), Polyommatus (Agrodiaetus) urmiaensis Schurian et ten Hagen, 2003, Polyommatus (Agrodiaetus) vanensissheljuzhkoi (Forster, 1960), Polyommatus (Agrodiaetus) vaspurakani (Lukhtanov et Dantchenko, 2003) and Polyommatus (Agrodiaetus) zarathustra Eckweiler, 1997.
Bayesian analysis was performed using the program MrBayes 3.2.2 (Ronquist et al. 2012 (link)). A GTR substitution model with gamma distributed rate variation across sites and a proportion of invariable sites was specified before running the program for 5,000,000 generations with default settings. The first 1250 trees (out of 5000) were discarded as a burn-in prior to computing a consensus phylogeny and posterior probabilities.

Plant material – We have extended the dataset of Potentilla species used in Eriksson et al. [2]
[3]  in order to make a more detailed study of the phylogenetic relationships within the genus and to identify major clades of related taxa. Our analysis includes 64 ingroup taxa and seven outgroup taxa from various genera in the sister clade Fragariinae. In this study, the taxonomy is based on Atlas Flora Europeae [16]  for the European species and the International Plant Name Index [33] for the non-European species. The aim has been to sample as many of the major groups proposed by Wolf [4] as possible. In addition, species belonging to the segregate genera Ivesia, Horkelia, Horkeliella Rydb. and Comarella Rydb. (the latter a section in Ivesia in recent treatments [34] ) have been included in the analysis. Attempts were also made to include Ivesia arizonica (Eastw. ex J.T.Howell) Ertter (first described as Purpusia saxosa Brandegee) and Ivesia santolinoides A. Gray (later transferred to Stellariopsis santalinoides (A.Gray) Rydb.) but this failed due to a lack of good material to extract DNA from. In total, eight representatives from these segregated North American genera were included. Material was collected in botanical gardens, from herbarium specimens or from natural populations (Table S1). Only seeds were available for some of the included taxa, and these were sown in the spring of 2006, and leaf material was collected for DNA extraction. Vouchers were prepared when plants were in flower (stored at GB herbarium, Gothenburg).
Morphological study – The results from previous studies imply that the style characters used by Wolf [4] may have taxonomic value [2]
[3] . Anther shape (Fig. 2) is also a character that has been proposed to carry information about species relationships, and has been used to subdivide Potentilleae [5] . Here we are interested in evaluating their usefulness within Potentilla as currently circumscribed by performing an ancestral state reconstruction analysis. The character states were determined by examining herbarium specimens and are summarized in Table S1. Ancestral character states were estimated by fitting three different models of rate variation, using maximum likelihood optimization and the nuclear data phylogeny. A likelihood-ratio test was used to choose which of the three models to apply. The first model assumes equal transition rates between all states (ER). In the second model, each pair of states can have a distinct rate of interchange, but this rate apply to both the forward and reverse transformation (SYM). The third model allows each pair of states to have distinct rates for both the forward and reverse transformation (ARD). The nuclear phylogeny was first converted to a fully dichotomous tree by adding zero length branches in the unresolved clades using the default setting of the function multi2di, and the analyses where then performed using the function ace in the package ape [35] of the statistical program R [36] .
Chromosome counts – Roots were pretreated in a 1:1 mixture of 0.3% Colchicin and 2mM 8-hydroxy-chinolin for 2 h at 15–20°C and transferred to Carnoy I (Acetic Acid and 95% Alcohol, 1:3) for fixation. The tissue was hydrolysed in a 1:1 mixture of concentrated Hydrochloric Acid and 95% Alcohol at room temperature for 5–10 min, washed twice with water, and stained in Aceto-Orcein for 0.5–1 h. The actively dividing root tip was dissected on a microscope slide and the cell mass squashed under a cover glass. To make slides permanent, the cover glass was removed after freezing in liquid Nitrogen. The tissue was washed and dehydrated in a series of 70%, 95% and absolute Alcohol. Before embedding in mountant the tissue was soaked in Histoclear for 24 h.
Extraction – Leaf material from wild or cultivated specimens was dried in silica gel before extraction. In some cases silica dried material was unavailable and herbarium material was used. For the majority of the samples, total genomic DNA was extracted using the DNeasy Plant Mini Kit (Qiagen, Valencia, California, USA). Standard protocol for plant tissue extractions was used. A minor part of the samples were extracted using the E.Z.N.A SP Plant DNA Miniprep kit (OMEGA Bio-Tec, Doraville, Georgia, USA) according to the enclosed protocol.
PCR-amplification – For the ITS and trnS/G regions we used PuReTaq Ready-To-Go PCR Beads [37] . Each reaction contained 1µl [20 µM] 5'-primer (forward) and 1µl [20 µM] 3'-primer (reverse), 0,5-4µl (typically 2µl) template DNA of unknown concentration and ultra purified water to a final volume of 25µl. Amplification of the ETS and trnL/F regions were performed using 25 µl MasterAmp 2x PCR PreMix G [38] , 1,5µl of [20 µM] forward and reverse primers, 0,2µl Termoprime Plus DNA Polymerase [39] , 0,5-4µl (typically 1µl) template DNA and ultra purified water to a final volume of 50µl. For a number of taxa extracted from herbarium material the MasterAmp 2x PCR PreMix G protocol produced low or no product at all. In these cases the PuReTaq Ready-To-Go PCR Bead protocol was used instead. PCR-products were purified with the QIAquick PCR Purification kit from QIAGEN, according to the enclosed protocol.
Sequencing – Dye terminator cycle sequencing with DTCS-Quick start kit (GenomeLab) were performed using a Beckman Coulter CEQ 8000 Genetic Analysis system (software v. 8.0) automated sequencer according to the manufacturer’s protocol.
Primers used for PCR amplification and sequencing - ETS1 and IGS6 [40] for the ETS region; ITS-1 [41] , ITS2 and ITS4 [42] and ITS3B [43] for the ITS region; trn-C, trn-D, trn-E and trn-F [44] for a trnL intron and the adjacent trnL-F spacer; trnS^GCU, 3´trnG^UUC, 5´trnG2G and 5´trnG2S [45] for a terminal intron in the trnG region and the spacer between trnS and trnG. All regions were sequenced in both directions using both the terminal PCR primers and two internal primers (except for the ETS region where no internal primers where used).
Sequence preparation - Sequences were assembled and manually edited using the Staden package version 1.6.0 [46] , using phred v.0.020425.c [47] for base calling and phrap v.0.990329 [48] for assembling contigs. Sequences were aligned with the software Muscle v. 3.6 [49] using the default settings followed by some additional manual editing in SeaView 4.2 [50] . Sequences from the chloroplast intergenic spacers trnL-trnF and trnS-trnG, were truncated in conserved regions of the 5' and 3' ends of the sequences and concatenated into a single matrix. Parts of the aligned matrix (corresponding to position 599-671 in sequence FN594698 from P. thuringiaca) around the internal primers were excluded due to sequencing problems of this region. The nuclear markers ITS and ETS were concatenated into a single matrix and truncated in both ends of the respective regions. Indels in both matrices were coded using SeqState v.1.32 [51] using the simple coding strategy [52] .
Test for recombination – The combined ETS and ITS dataset was analyzed for indications of recombination with the following methods: RDP [53] ; Bootscan/Recscan [54] ; Geneconv [55] ; MaxChi [56] ; Chimera [57] ; SiScan [58] and 3Seq [59] , implemented in the RDP3 program v.3.34 [60] . Cut off of P-values where set to 0.05 with a Bonferroni correction, otherwise using the default settings. Putative recombination events that were detected by two or more methods were further investigated with the Bootscan method [54] , using 100 bootstrap samples and a window size of 100-300 positions.
Phylogenetic analysis - Both datasets were analyzed with MrBayes v.3.2 (source code accessed with cvs 22 January 2009) compiled to use the MPI parallel library [61]
[62] . Two independent analyses where run for 2 000 000 generations (nuclear data set) or 5 000 000 generations (chloroplast data set), with eight chains each and the Temp parameter set to 0.1. Trees were sampled every 1000 generations. The first 25% of the sampled trees were discarded and the remaining samples were summarized in a 50% majority-rule consensus tree. Both the nuclear and the chloroplast matrices were analyzed under the general time-reversible model (GTR) assuming a gamma shaped distribution of rate variation. MrAic.pl v1.4.3 [63] in conjunction with PHYML v.2.4.4 [64] was used to select the model. All three information criterions implemented in MrAic (AIC, AICc and BIC) rated this evolutionary model the highest. A binary model was used for the coded gaps. The mixing behavior of the mcmc chains was analyzed in Tracer v1.5 [65] and was found to have been sufficient. Mixing between the different metropolis coupled chains were analyzed in the statistics package R [36] by plotting selected columns in the *.mcmc file. In addition, two more analyses were performed based on the two matrices using the same method, but excluding the gap codes (data not shown). 

After maturation, COCs were denuded by incubation with hyaluronidase solution for 10 min at 37 °C and then vortexed for 5 min. COCs were fixed in an ethanol-acetic acid solution (three parts ethanol and one part acetic acid) for 24 h and stained with 1% orcein solution (1 g orcein in 100 mL ethanol-acetic acid solution). Nuclear maturation stage was evaluated under a phase contrast microscope (Axiostar plus, Carl Zeiss, Jena, Germany) using a contrast phase filter N° 2 at 400× magnification. Oocytes were classified as germinal vesicle (GV), germinal vesicle breakdown (GVBD), metaphase I (MI), or metaphase II (MII). Nuclear maturation was calculated by the number of oocytes that achieved metaphase II (MII) stage divided by the total number of evaluated oocytes.