Skip to main content
Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Unlocking the Karyological and Cytogenetic Diversity of Iris from Lebanon: Oncocyclus Section Shows a Distinctive Profile and Relative Stasis during Its Continental Radiation

  • Nour Abdel Samad,

    Affiliations Faculté des Sciences, Département Sciences de la Vie et de la Terre, Laboratoire Caractérisation Génomique des Plantes, Campus Sciences et Technologies, Université Saint-Joseph, Mar Roukos Mkalles, Lebanon, Ecologie, Systématique, Evolution, UMR 8079 Univ. Paris-Sud, AgroParisTech, Université Paris-Saclay, Bat. 360, 91405 Orsay, France

  • Magda Bou Dagher-Kharrat ,

    Affiliation Faculté des Sciences, Département Sciences de la Vie et de la Terre, Laboratoire Caractérisation Génomique des Plantes, Campus Sciences et Technologies, Université Saint-Joseph, Mar Roukos Mkalles, Lebanon

  • Oriane Hidalgo,

    Affiliation Royal Botanic Gardens, Kew, Richmond, Surrey, TW9 3AB, United Kingdom

  • Rana El Zein,

    Affiliation Faculté des Sciences, Département Sciences de la Vie et de la Terre, Laboratoire Caractérisation Génomique des Plantes, Campus Sciences et Technologies, Université Saint-Joseph, Mar Roukos Mkalles, Lebanon

  • Bouchra Douaihy,

    Current address: Faculté des Sciences, Département de Biologie, Université Libanaise, Tripoli, Lebanon

    Affiliation Faculté des Sciences, Département Sciences de la Vie et de la Terre, Laboratoire Caractérisation Génomique des Plantes, Campus Sciences et Technologies, Université Saint-Joseph, Mar Roukos Mkalles, Lebanon

  • Sonja Siljak-Yakovlev

    Affiliation Ecologie, Systématique, Evolution, UMR 8079 Univ. Paris-Sud, AgroParisTech, Université Paris-Saclay, Bat. 360, 91405 Orsay, France


Despite being an important target of conservation concern and horticultural interest, Lebanese irises yet have a confusing taxonomic history and species’ delimitation is often considered problematic, more especially among royal irises (Iris section Oncocyclus). Indeed, these irises of exceptionally large and spectacular flowers have radiated across Caucasus and eastern Mediterranean giving rise to a number of strict endemic taxa, many of them being considered under threat. Whilst efforts have mostly focused on clarifying the evolutionary relationships in the group based on morphological and molecular data, karyological and cytogenetic characters have been comparatively overlooked. In this study, we established for the first time the physical mapping of 35S rDNA loci and heterochromatin, and obtained karyo-morphological data for ten Lebanese Iris species belonging to four sections (Iris, Limniris, Oncocyclus and Scorpiris). Our results evidenced distinctive genomic profiles for each one of the sections, where Oncocyclus irises, while having the lowest chromosome numbers, exhibit both the highest number of 35S loci and CMA3+ sites. The continental radiation of royal irises has been accompanied by a relative karyological and cytogenetic stasis, even though some changes were observed regarding karyotype formula and asymmetry indexes. In addition to that, our results enabled taxonomic differentiation between I. germanica and I. mesopotamica–two taxa currently considered as synonyms–and highlighted the need for further studies on populations of I. persica and I. wallasiae in the Eastern Mediterranean Region.


The genus Iris L. includes about 280 species distributed across the temperate region of the Northern Hemisphere [1]. Recurrent hybridization, which has given rise to a myriad of garden forms [2] has also largely contributed to make this genus the largest and most complex of Iridaceae [3]. The classification of Iris is indeed a difficult task to tackle; botanists and taxonomists are still far from reaching a consensus on this issue. This problem is certainly reflected through the different subgeneric and sectional classifications established on the basis of morpho-anatomical features, ecological and cytogenetic traits [4,5,6,7]. The most recent taxonomic revision [5] recognizes six subgenera: Nepalensis Dykes, Xiphium (Miller) Spach, Scorpiris Spach, Hermodactyloides Spach, Iris L. and Limniris Tausch. According to Mouterde [8], only the four latter subgenera occur in Lebanon. The subgenus Iris is represented by two sections: Iris L. with two taxa and Oncocyclus (Siemssen) Baker with seven taxa, while the subgenus Limniris is represented by two taxa, the subgenus Hermodactyloides with one taxon and subgenus Scorpiris with two taxa.

This study focused on Iris section Oncocyclus–the royal irises–which includes numerous strict endemic species. These irises are native to the Near East, especially the Caucasus region, eastern Turkey, Syria, Lebanon and Israel and extend to the Negev Desert. In the east they are found in Iraq, Iran and Afghanistan [9]. Oncocyclus section includes 33 species (up to 45 taxa considering subspecies, forms and varieties), which are all regional endemics [10]. In the eastern Mediterranean region, they grow in disjunctive populations separated by short geographical distances [11]. In a recent study, Wilson et al. (2016) [12] suggested the Caucasus as the ancestral area for Oncocyclus section and the Eastern Mediterranean region as an important area of diversification. From a nomenclature and systematic point of view, Oncocyclus section is particularly challenging within Iridaceae [5] and species concept has already been debated [13]. Taxonomic circumscription has been based on minor differences in plant size, leaf shape and flower color. It appears that many local forms have been considered as species, which increased the number of taxa. In addition, the sympatric distribution of some species and interfertility among Oncocyclus species [14] could have led to a large number of hybrids, often recognized as full species.

Over the years, taxonomic treatment of the section Oncocyclus has experienced a ‘lumping’ change. From 65 species considered by Avishai in 1977 [15], Mathiew in 1989 [5] proposed 41 species and in 1997 Rix [10] proposed only 30. Some sympatric species were recognized as hybrids, but the grouping trend affected mainly the Caucasian species, and no complete or inclusive taxonomic revision of Oncocyclus section in the Levant has been carried out to date.

The Lebanese Oncocyclus irises are represented by seven taxa (Fig 1) of which four are considered to be strict endemics [11]: I. cedreti Dinsmore ex Chaudhary, I. sofarana Foster subsp. kasruwana (Dins.) Chaudhray, I. sofarana subsp. sofarana and I. westii Dinsmore. They occur in distinct populations across the Mount Lebanon chain [10] between 1200 and 2000 m of altitude. The three other Lebanese Oncocylus species are I. bismarckiana Regel (Lebanon, Syria and Israel) and I. lortetii Barbey ex Boiss. (Lebanon and Israel) and I. antilibanotica Dinsm. (Lebanon and Syria).

Fig 1. Diversity of Lebanese royal irises, illustrated by some of the studied taxa.

A: Iris sofarana subsp. sofarana, B: I. sofarana subsp. kasruwana, C: I. cedreti, D: I. westii, E: I. bismarckiana, F: I. lortetii, G: I. antilibanotica. Photographs from M. Bou Dagher-Kharrat

The botanical descriptions of the endemic Lebanese Oncocyclus species, that have been written more than 30 years ago on the basis of a limited sample size, provide however a good illustration of the variability found in populations nowadays [16]. Oncocyclus irises are morphologically very close and may be confused because they all have different variants with similar colors, more or less dark purple [5,8,9,17,18]. It is worth noting that the differentiation between I. sofarana subsp. sofarana, I. sofarana subsp. kasruwana and I. westii according to Mouterde, is only based on the size of the individuals and the shade of standards color.

From a conservation point of view, Iris genus includes 53 taxa that are threatened worldwide, of which 29 belong to the Oncocyclus section [19]. Oncocyclus irises are narrow endemic and of high conservation priority in Lebanon [20,21]. These plants with conspicuous flowers are threatened by intensive collection and habitat destruction. In Lebanon, conservation efforts are now undertaken in order to conserve Iris populations through legislation and establishment of micro-reserves [22]. Based on their complete self-incompatibility, fertilization of Oncocyclus irises relies on pollination by night-sheltering solitary male bees [23,24,25,26] and Honey bees as frequent diurnal visitors [27]. Nevertheless, pollination is dramatically hindered by the intensive use of pesticides on the adjacent agriculture lands (Bou Dagher-Kharrat, personal observation). Even though they show a dense clonal growth, populations can be sometimes restricted to few dozens of individuals.

Rescue from possible extinction could be performed through the reinforcement of populations by introducing individuals from adjacent populations. In this regard, defining species and differentiating taxa is a crucial and a prerequisite in order to define conservation units. Indeed, conservation strategies require knowing which units (species, subspecies, or populations) need to be preserved and how unique they are.

Wilson et al. [12] used in 2016 plastid and nuclear DNA sequences to infer Oncocyclus phylogenetic relationships. They found that the species diversified in isolation especially in the Eastern Mediterranean region where populations are scattered on the mountains across rocky hillsides and steppes. However, continental radiation of the royal irises did not involve necessarily chromosome rearrangements, since chromosome number (2n = 20) and their karyotype feature remained basically unchanged throughout Oncocyclus taxa [15]. Could molecular cytogenetic bring more insight into this complex Iris group?

The physical mapping of rDNA and the distribution of heterochromatin turn out to be a source of chromosomal markers for identifying several chromosome regions [28,29,30,31,32,33] and provides according to Dobigny et al. “valuable information on homologies between chromosomal segments, mainly between closely related species” [34,35,36]. In addition, this set of markers may constitute a phylogenetic tool to detect genome evolution during speciation [37,38,39]. Molecular cytogenetics allowed indeed to detect interesting variation in the rDNA markers as revealed by fluorescent in situ hybridization (FISH) patterns in Iris subgenus Xiphium and evidenced a species-dependent pattern of rDNA sites [40]. However, such studies on the Iris genus remain scarce.

The present study combines classical karyological approach and physical mapping of heterochromatin and rRNA genes in order to characterize the genome organization of ten Lebanese Iris taxa. The main objective of this study was to contribute to the understanding of phylogenetic relationships, more particularly for the species of Oncocyclus section, by checking possible cytogenetic differences between the investigated taxa. For this purpose, besides classical karyological approach, karyotypes were characterized using molecular cytogenetic techniques: FISH for physical mapping of 18S-5.8S-26S (35S) and 5S rRNA genes and fluorochrome banding for distribution patterns of GC-rich heterochromatin regions.

Material and Methods

Plant material

In order to obtain fresh root meristems for chromosome preparations, rhizomes of ten Iris taxa were collected from ten Lebanese natural populations. Their geographical origins are presented in Table 1.

Table 1. Geographical origin (locality and altitude) and flowering date of the investigated taxa.

Rhizomes were then potted in perlite.

We followed the nomenclature of Mouterde published in 1983 [41], although its correspondence with recent nomenclature [42] is also provided in S1 Table. No specific permissions were required for populations located on public lands. Permissions are obtained from religious communities and private land owners for the other populations. Irises are not yet protected in Lebanon. Herbarium specimens are deposited in the National Herbarium at the Lebanese University of Beirut.

Chromosome preparation and construction of karyotypes

Root tips obtained from rhizomes were pre-treated 24h in 0.05% Colchicine at 4°C or 5h at room temperature. The fixation was performed in Carnoy’s solution (3:1 absolute ethanol: glacial acetic acid) at 4°C for at least 24–48h.

For morphometrical karyotype analysis, the meristems were hydrolyzed in 1 N HCl at 60°C for 12 min, stained in and squashed in 2% (w/v) aceto-orcein. Slides were freezed at -80°C during 24 h, then cover slips were removed, preparations were dried at least 24 h and then mounted in Euparal. Chromosome counts were made on well-spread metaphase plates. The karyotype was determined by examining five metaphase plates per species. Determination of centromere position and chromosome type were made according to Levan & al. (1964) [43]. The following characters were measured: long arm (l); short arm (s); total chromosome length (TL);; arms ratio (r = ratio long/short arms); centromeric index (Ci% = 100 x s/TL);; chromosome type (Ct; according to Levan & al., 1964) [43]; mean centromeric asymmetry (MCA = Ax100 according to Peruzzi & Eroğlu, 2013) [44]; A = Mean(long arm-short arm)/(long arm+short arm) according to Watanabe & al. (1999) [45]; coefficient of variation of chromosome length CVCL = A2 x 100 (Paszko, 2006) [46] where A2 = standard deviation of chromosome length/mean chromosome length (Romero Zarco, 1986) [47];

Chromosome preparation for fluorochrome banding and FISH

Chromosome plates for fluorochrome banding and the FISH experiment were prepared using the air-drying technique of [48], with slight modifications. Root tips were washed in a citrate buffer (pH 4.6) for 10 min and then transferred into the enzyme mixture [4% R-10 cellulase (Yakult Honsha Co. Tokyo, Japan), 1% pectolyase Y-23 (Seishin Co. Tokyo, Japan), 4% hemicellulase (Sigma)] in a moist chamber at 37°C for 15 min. The digested meristems were gently squashed in a drop of acetic-acid 45%. Cover slips were removed after freezing over night at -80°C.

For detection of GC-rich DNA regions, chromomycin A3 banding (CMA, Sigma) was performed following the technique of [49] with minor modifications [50]: slides were stained with 0.2 μg/ml of CMA solution for 1 h in the dark and mounted in Citifluor AF1 anti-fade agent (Agar Ltd). Some slides stained with chromomycin were destained in fixative (3:1 absolute ethanol:glacial acetic acid), dehydrated in a graded ethanol series (70%, 90%, 100%), air-dried for at least 12 h at room temperature, and then used for the FISH experiment.

A double FISH experiment was carried out with two DNA probes. The 35S rDNA probe was a 4 kb clone from the EcoRI fragment, including 18S-5.8S-26S rDNA sequences from Arabidopsis thaliana labeled with direct Cy3 fluorochrome (Amersham, Courtaboeuf, France) by nick translation, according to the manufacturer’s protocol. The 5S rDNA probe was the pTa794 clone [51] containing a complete 410 bp BamHI fragment of wheat, including the gene (120 bp) and the spacer (290 bp). The probe was labeled with digoxigenin-11-dUTP (Roche Diagnostics, Meylan, France) after polymerase chain reaction (PCR) using universal M13 primers, and antibody detection was made with anti-digoxigenin-fluorescein (Roche Diagnostics GmbH). The probe mixture consisted of 1–2 ng/μl of each probe, 50% (v/v) formamide, 10% (w/v) dextran-sulfate, 0.1% (w/v) sodium dodecyl sulfate, 250 μg/ml salmon sperm, 20× sodium saline citrate (SSC), completed with ultrapure water. In situ hybridization was carried out following the method of [52]. Slides were counterstained and mounted in Vectashield medium containing DAPI (4′, 6-diamidino-2-phenylindole, Vector Laboratories). For rDNA site distribution analyzes, a minimum of 10 well-spread metaphase plates were analyzed for each Iris species.

Microscopy and chromosome analysis

Chromosome observations were performed using an Epifluorescence Zeiss Axiophot microscope with different combinations of excitation and emission filter sets (01, 07, 15 and triple filter set 25). The signals were analyzed using the highly sensitive CCD camera (RETIGA 2000R; Princeton Instruments, Every, France) and an image analyzer (Metavue, Every, France).


Chromosome number and karyotype analysis

A diploid chromosome number of 2n = 2x = 20 with basic chromosome number x = 10 was observed in the seven studied Oncocyclus taxa. Their detailed karyotypes were established for the first time in this study. Morphometric data for each karyotype are presented in S2 Table. The karyotypes of the Oncocyclus taxa were quite similar in size and symmetry (S1 Fig). However, small differences are perceptible at closer look. I. lortetti for instance is the only taxa among the Oncocyclus analyzed to have (2t+18st). Chromosome lengths varied from 1.8 to 6.7 μm.

The remaining non Oncocyclus taxa presented cytotypes with different chromosome numbers and ploidy levels: diploid 2n = 2x = 24 for I. persica, tetraploid 2n = 4x = 40 for I. unguicularis var. cretensis and 2n = 4x = 48 for I. mesopotamica with basic chromosome numbers of x = 12 and x = 10 respectively. Morphometric data are represented in Table 2.

Table 2. Main data on karyotype features of investigated Iris taxa.

Total chromosome length of haploid complement varied from 30.5 to 44.5 μm in Oncocyclus irises while karyotype symmetry index varied between 84.2% and 86.8%. Total chromosome length and the karyotype symmetry index were respectively 81.2 μm and 60.4% for I. persica, 74.89 μm and 63.57% for I. unguicularis var. cretensis and 86.7 μm and 43% for I. mesopotamica.

The mean centromeric asymmetry (MCA) of Oncocyclus taxa varied from 68.4 to 73.6%. For I. persica, I. unguicularis var. cretensis and I. mesopotamica, MCA were 40.4%, 27.13% and 39.9% respectively. The CVCL of the ten taxa, ranged from 20.9% to 33% (Table 2).

Heterochromatin and ribosomal genes mapping

The physical mapping of two rRNA gene families 5S and 35S, and the distributional pattern of GC-rich DNA regions (heterochromatin) in the chromosomes of ten Lebanese taxa have been performed for the first time in this study. The results are presented in Table 3 and Fig 2.

Fig 2.

Metaphase chromosome plates and Idiograms of Iris taxa AK: Metaphase chromosome plates of Iris taxa after double target FISH with 5S (green signals) and 18-26S rDNA (red signals) probes. C1 represents CMA staining (yellow signals). A’K’: Idiograms with location of 5S (green) and 18-26S rDNA (red) rRNA genes. A’: I. sofarana subsp. sofarana (Falougha), B’: I. sofarana subsp. sofarana (Hazzerta) C’: I. sofarana subsp. kasruwana, D’: I. cedreti, E’: I. westii, F’: I. bismarckiana, G’: I. lortetii, H’: I. antilibanotica, I’: I. persica, J’: I. unguicularis var. cretensis, K’: I. mesopotamica. Scale bar 10 μm.

Table 3. Synthesis of results concerning molecular cytogenetic approach of investigated Iris taxa.

Fluorochrome banding revealed GC-rich heterochromatin regions (CMA+ bands) which were always associated with 35S rDNA loci. Consequently, the number and position of GC-rich DNA regions correspond to those of 35S loci (Table 3). G-C rich heterochromatin and rDNA organization was conserved and almost identical for all Oncocyclus taxa (Fig 2A–2H). The 35S rRNA genes (red signals) were situated in the secondary constrictions (SC) and satellites, which correspond to Nuclear Organized Regions (NORs) of chromosome pair 5 and 6 and on the short arm of chromosome pair 7. Four nucleolar silver-stained granules of active nucleolus organizer regions in interphase cells of I. sofarana and on I. mesopotamica were obtained confirming that all the 35S rRNA genes detected by FISH were active in I. mesopotamica while just 4 out of 6 were active in I. sofarana (Data not shown). This experiment should be conducted on the other species to confirm this tendency.

In I. persica ribosomal genes 35S were situated on the short arms of chromosome pairs 11 and 12 (Fig 2I and 2I’), in I. unguicularis var. cretensis on the pairs 19 and 20 (Fig 2J and 2J’), and in I. mesopotamica on the pairs 6 and 11 (Fig 2K and 2K’). The only 5S locus (green signals) was located intercalary on paracentromeric position of telomeric chromosome pair 9 in all Oncocyclus taxa (Fig 2A–2H), on chromosome pair 6 in I. persica, on pair 8 in I. unguicularis var. cretensis and on pair 17 in I. mesopotamica (Fig 2I–2K).


Chromosome number and karyotype features

The karyotypes of investigated taxa were established for the first time using conventional measurements of chromosomes on several metaphase plates per species or population.

Our results show that the karyotype of Oncocyclus species remained mainly unchanged. The species showed an extremely similar, bimodal, asymmetric chromosome complement (2n = 20) with four pairs (or five in I. lortetii) of long chromosomes and six pairs of small ones (Fig 2). Among the long chromosomes, two pairs with a “conspicuous knoblike” short arm and two pairs with “minute”, almost indistinct short arms were observed. The small chromosomes could be differentiated into two satellite pairs, two pairs with knoblike short arms, and two other pairs with minute short arms. All the species from Oncocyclus section studied here were found to be diploid with basic chromosome number x = 10 and to present a very close karyotype profile. This is in accordance with results reported by [15,55] for a dozen of Oncocyclus taxa analyzed. The constant chromosome number and similar karyotype feature should be considered as a characteristic of the entire section and a diagnostic trait separating the Oncocylus from other Iris sections.

In addition to the basic chromosome number of x = 10 found in the Oncocyclus and Limniris sections of the subgenus Iris, x = 12 was found in I. mesopotamica (Iris section) and I. persica (subgenus Scorpiris). Basic chromosome numbers of x = 8, 10, 11 and 12 were reported in subgenus Iris and x = 10, 11, 12 and 13 in subgenus Scorpiris [56]. When comparing basic chromosome number with phylogenetic inferences of Iris genus [57], we did not find an explicit trend or correlation between basic chromosomes number and evolutionary history. In each subgenus, different basic chromosome numbers exists. Therefore, we cannot assume neither the smallest nor the biggest basic chromosome number is attributed to the ancestral number.

It is noteworthy that I. persica taxonomic status in Lebanon is still debated. In fact, Hall & Seisums [58] considered some Iris persica populations of Lebanon and Syria as a new species called Iris wallisiae. They contradicted Mouterde (1966) who included I. persica in his treatment of the flora of Syria and Lebanon and declared that there is no evidence of I. persica ever having been found in Lebanon. Furthermore, according to Hall & Seisums (2014) [58] all reports of I. persica by Tohmé & Tohmé [59,60,61] for Bekaa valley refer to I. wallisiae. A chromosome count of 2n = 22 was found by Hall and Seisums (2014) in individuals from three Syrian populations of I. wallisiae whereas it was 2n = 20 in six Syrian populations of I. persica. Unexpectedly, our results differ from these chromosome counts, since we found 2n = 24 in the presumed I. persica population analyzed. Further studies carried out on a larger number of populations in this geographical area need to be conducted to better understand the taxonomy and phytogeography of these taxa.

For I. mesopotamica we found 2n = 4x = 48 which is in agreement with earlier reports by [62,63]. Iris mesopotamica is considered a synonym of I. germanica [42]. Iris germanica was found by Siljak-Yakovlev [64] to be also tetraploid. Both are considered as hybrids and found only in cultivated areas or in the wild having escaped from cultivation. Iris mesopotamica is propagated vegetatively and distributed locally, giving the impression of having a stable population with a distinct geographical range [5]. Frequently grown in cemeteries and gardens, it is supposed to come from the Asian Mediterranean coasts (Mouterde, 1983).

For I. unguicularis var. cretensis, we confirm the previous report of 2n = 4x = 40 [65]. This variety is also found in Greece, Crete and Minor Asia. Typical forms of I. unguicularis are widespread in the Eastern Mediterranean region and North Africa where they display a great variability.

Polyploidy was clearly important in the early diversification of the Iridaceae [66]. In Iris, where changes in basic chromosome numbers are common, the ancestral base number, however, remains unclear [66]. The phylogenetic reconstructions do not provide an adequate resolution to interpret neither an increasing nor a decreasing (descending) dysploidy. Although, few studies evaluate dysploidy variation of chromosome number in a phylogenetic framework, a decrease in chromosome numbers comes out not to be unusual [67,68,69,70]. Decreasing dysploidy has been already proposed for several genera of the Iridaceae [66].

Based on the study of the karyotypes feature, we can conclude that Oncocyclus is more recent than Iris, Limniris and Scorpiris sections. The constant chromosome number (2n = 20) and extreme similarity of karyotypes among the species of Oncocyclus section can be related to its small geographical distribution and shows that Oncocyclus is a recent–still evolving–section.

Distribution pattern of GC-rich DNA regions and rRNA genes

The comparative chromosome mapping of rRNA genes and GC-rich DNA regions have been established here for the first time for all ten investigated taxa. The rDNA is known to participate actively in genome rearrangements [71,72], thus analyzing and comparing 35S and 5S rDNA profiles across related species potentially allow to test hypotheses on their relationships.

Although this kind of studies remains very scarce for the Iris genus, the chromosomal locations of 5S and 35S rDNA loci have been previously determined for seven species of subgenus Xiphium [40], with a possible trend of increased 35S rDNA loci number during the diversification of the group.

In our study, the two rRNA gene families were located on separate chromosome pairs; thus, the 5S genes are of S-type arrangement [73]. Six signals of 35S rDNA and two signals of 5S rDNA are observed in diploid Oncocyclus irises while four signals of 35S and two signals of 5S were detected in tetraploid I. unguicularis var. cretenis and I. mesopotamica. Polyploidy is frequently associated with epigenetic silencing of rDNA loci [74,75]. Those units and loci that are inactive could be most vulnerable to deletion since their loss would have no selective consequence [76]. In Iris germanica which is currently considered as synonym of I. mesopotamica, Siljak-Yakovlev [64] found eight loci of 35S and two loci of 5S. This different 35S rDNA signals number between these two species questions their real synonymy and call for further studies. Both are tetraploid but their presumed hybrid origin should be confirmed.

Our results prove once again what was reported in previous studies: a loss in 35S and perhaps 5S rDNA loci can be considered as an early indication of genome diploidization in polyploid. In allopolyploids, various types of genetic alterations are observed as reduction in copy number, locus loss, intra- and intergenomic recombination [77,78,79,80]. It was the case for I. versicolor, an allopolyploid hybrid for which 18–26S rDNA is likely to have undergone a locus loss rather than locus co-evolution [76]. If this should be applied to the polyploidy of I. germanica and I. mesopotamica, the first one could be an autopolyploid while the other one could be an interspecific hybrid. Of course, the polyploidy origin should be addressed more carefully by including in the study the putative parents. At this point we can only confirm that ribosomal RNA genes mapping enables the distinction between I. germanica from Balkans and I. mesopotamica.

Outlandishly, this technique failed to show differences between the seven Oncocyclus irises studied. This finding supports their recent common evolutionary history, as previously suggested [18] Although molecular markers found the two I. sofarana subspecies as polyphyletic [12], interbreeding between the Oncocyclus species is reported to be possible, and creates vital progenies, with fertile hybrids [11]. However, several factors limiting gene flow between populations could have conduced to speciation and selection of the locally adapted genotypes: indeed, (i) the different populations of the Lebanese taxa analyzed in this study are highly fragmented and not close enough for allowing their short flying distance pollinators to maintain gene flow; furthermore, (ii) these taxa are found at different altitudes and flowering dates do not overlap thus contributing to their genetic isolation; finally, (iii) Oncocyclus irises are myrmecochory plants, and having their seeds dispersed by ants could also contribute to hinder gene flow. For conservation issues, when population reinforcement will be considered as an option, it is advisable to realize controlled cross-fertilization experiments before in order to confirm their biological compatibility. This kind of information is crucial in order to address IUCN red list status of these taxa.

Concluding Remarks

This is a contribution to the knowledge of this large genus which has been, until now, rarely studied in this area of research. It provides a first karyomorphological and molecular cytogenetic characterization of ten Lebanese Iris species. An important inter-sections variation was detected among the studied taxa. Difference in chromosome number, morphometric data of karyotypes, number and position of GC rich DNA regions and 5S and 35S rDNA loci were evidenced among three species belonging to sections Iris, Limniris and Scorpiris. However the absence of variability and the strong similarity in the karyotype features observed among Oncocyclus taxa suggest that this is probably a very young group whose speciation is still in progress as reported in a recent phylogeny including some Oncocyclus representative [12]. Additional molecular and cytological studies, including a wider sampling of Oncocyclus irises from the eastern Mediterranean, are currently underway. These studies should clearly describe the diversification and speciation of this section.

Supporting Information

S1 Fig. Metaphase plates and karyotypes of selected Iris taxa stained with acetoorcein


S1 Table. Accepted names of studied irises and their synonyms according to The Plant List, 2013


S2 Table. Morphometric data concerning the karyotype of Iris taxa



We thank Odile Robin and Fatima Pustahija for their technical assistance in molecular cytogenetics (Ecologie, Systématique, Evolution, UMR 8079, Université Paris Sud), Dr. Georges Tohmé and Mr. Nasser Chreif for their help in collecting some of the plant material.

Author Contributions

  1. Conceptualization: MBK SSY.
  2. Data curation: SSY NAS.
  3. Formal analysis: NAS OH.
  4. Funding acquisition: MBK.
  5. Investigation: BD REZ NAS.
  6. Methodology: MBK SSY.
  7. Project administration: MBK.
  8. Resources: BD REZ NAS.
  9. Software: OH.
  10. Supervision: MBK SSY.
  11. Validation: MBK SSY.
  12. Visualization: MBK.
  13. Writing - original draft: MBK SSY NAS.
  14. Writing - review & editing: MBK.


  1. 1. Rasoul TN (1984) Ornamental bulbs. Mosul: Al-Mousel University.
  2. 2. Glasgow K (1997) Irises. A practical gardening guide. Portland, Oregon: Timber Press.
  3. 3. Makarevitch I, Golovnina K, Scherbik S, Blinov A (2003) Phylogenetic Relationships of the Siberian Iris Species Inferred from Noncoding Chloroplast DNA Sequences. Int J Plant Sci 164: 229–237.
  4. 4. Doronkin VM (1987) Iridaceae Juss. Nauka, Novosibirsk. 113–125 p.
  5. 5. Mathew B (1989) The Iris. London: Batsford.
  6. 6. Rodionenko GI (1987) The genus Iris L.: questions of morphology, biology, evolution, and systematics. London: British Iris Society.
  7. 7. Wu Q-G, Cutler DF (1985) Taxonomic, evolutionary and ecological implications of the leaf anatomy of rhizomatous Iris species. Bot J Linn Soc 90: 253–303.
  8. 8. Mouterde P (1966–1983) Nouvelle flore du Liban et de la Syrie. Beyrouth (Liban): Imprimerie catholique.
  9. 9. Köhlein F (1987) Iris. Portland, Oregon: Timber Press.
  10. 10. Rix M (1997) Section Oncocyclus. In: A guide to Species Irises. London: Cambridge Press.
  11. 11. Avishai M, Zohary D (1980) Genetic Affinities among the Oncocyclus irises. Botanical Gazette 141: 107–115.
  12. 12. Wilson CA, Padiernos, J. & Sapir Y. (2016) The royal irises (Iris subg. Iris sect. Oncocyclus): Plastid and low-copy nuclear data contribute to an understanding of their phylogenetic relationships. Taxon: 12 pp.
  13. 13. Sapir Y, Shmida AVI (2002) Species concepts and ecogeographical divergence of Oncocyclus irises. Isr J Plant Sci 50: 119–127.
  14. 14. Avishai M (1977) Species relationships and cytogenetic affinities in section Oncocyclus of the genus Iris. Jerusalem: Hebrew University.
  15. 15. Avishai M, Zohary D (1977) Chromosomes in the Oncocyclus Irises. Botanical Gazette 138: 502–511.
  16. 16. West WA (1967) Iris antilibanotica. ASI Yearbook: 64–66.
  17. 17. Chaudhary SA (1972) Three new taxa of Iris subgenus Oncocyclus from Lebanon and Syria. Bot Notiser 125: 497–500.
  18. 18. Saad L (2006) Etude écogéographique et biologie de la conservation des Iris Oncocycles endémiques du Liban [PhD thesis]. Gembloux: Faculté universitaire des sciences agronomiques de Gembloux.
  19. 19. Walter KS, Gillett HJ (1998) 1997 IUCN Red List of threatened plants: IUCN.
  20. 20. Saad L, Mahy G (2009) Molecular and morphological variation of rare endemic oncocyclus irises (Iridaceae) of Lebanon. Bot J Linn Soc 159: 123–135.
  21. 21. Sapir Y, Shmida A., and Fragman O. (2003) Constructing Red Numbers for endangered plant species: Israeli flora as a test case. J Nat Conserv 11 91–107.
  22. 22. Bou Dagher-Kharrat M (2013–2016) Determination of Important Areas for Plants and Creation of Micro-Reserves to Conserve Rare or Endemic Species in Lebanon. In: Fund CEP, editor.
  23. 23. Lavi R, Sapir Y (2015) Are pollinators the agents of selection for the extreme large size and dark color in Oncocyclus irises? New Phytol 205: 369–377. pmid:25157604
  24. 24. Monty A, Saad L, Gg Mahy (2006) Bimodal pollination system in rare endemic Oncocyclus irises (Iridaceae) of Lebanon. Canadian Journal of Botany 84: 1327–1338.
  25. 25. Sapir Y, Shmida A, Ne’eman G (2005) Pollination of Oncocyclus irises (Iris: Iridaceae) by Night-Sheltering Male Bees. Plant Biol 7: 417–424. pmid:16025415
  26. 26. Vereecken NJ, Dorchin A, Dafni A, Hötling S, Schulz S, Watts S (2013) A pollinators' eye view of a shelter mimicry system. Ann Bot 111: 1155–1165. pmid:23599249
  27. 27. Watts S, Sapir Y, Segal B, Dafni A (2013) The endangered Iris atropurpurea (Iridaceae) in Israel: honey-bees, night-sheltering male bees and female solitary bees as pollinators. Ann Bot 111: 395–407. pmid:23275630
  28. 28. Adams SP, Leitch IJ, Bennett MD, Chase MW, Leitch AR (2000) Ribosomal DNA Evolution and Phylogeny in Aloe (Asphodelaceae). Am J Bot 87: 1578–1583. pmid:11080107
  29. 29. Bareka P, Siljak-Yakovlev S, Kamari G (2012) Molecular cytogenetics of Bellevalia (Hyacinthaceae) species occurring in Greece. Plant Syst Evol 298: 421–430.
  30. 30. Garnatje T, Hidalgo O, Vitales D, Pellicer J, Vallès J, Robin O, et al. (2012) Swarm of terminal 35S in Cheirolophus (Asteraceae, Centaureinae). Genome 55: 529–535. pmid:22794166
  31. 31. Jang T-S, Emadzade K, Parker J, Temsch EM, Leitch AR, Speta F, et al. (2013) Chromosomal diversification and karyotype evolution of diploids in the cytologically diverse genus Prospero (Hyacinthaceae). BMC Evol Biol 13: 1.
  32. 32. Ruas CdF Vanzela ALL, Santos MO Fregonezi JN, Ruas PM Matzenbacher N, et al. (2005) Chromosomal organization and phylogenetic relationships in Hypochaeris species (Asteraceae) from Brazil. Genet Mol Biol 28: 129–139.
  33. 33. Souza G, Crosa O, Guerra M (2015) Karyological, morphological, and phylogenetic diversification in Leucocoryne Lindl (Allioideae, Amaryllidaceae). Plant Syst Evol 301: 2013–2023.
  34. 34. Bogunić F, Siljak‐Yakovlev S, Muratović E, Ballian D (2011) Different karyotype patterns among allopatric Pinus nigra (Pinaceae) populations revealed by molecular cytogenetics. Plant Biol 13: 194–200. pmid:21143741
  35. 35. Dobigny G DJ-F, Robinson TJ, Volobouev V (2004) Cytogenetics and cladistics. Syst Biol 53: 470–484. pmid:15503674
  36. 36. Muratović E, Hidalgo O, Garnatje T, Siljak-Yakovlev S (2010) Molecular phylogeny and genome size in European lilies (Genus Lilium, Liliaceae). Advanced Science Letters 3: 180–189.
  37. 37. Cuadrado A, Schwarzacher T, Jouve N (2000) Identification of different chromatin classes in wheat using in situ hybridization with simple sequence repeat oligonucleotides. Theor Appl Genet 101: 711–717.
  38. 38. Cuadrado A, Vitellozzi F, Jouve N, Ceoloni C (1997) Fluorescence in situ hybridization with multiple repeated DNA probes applied to the analysis of wheat-rye chromosome pairing. Theor Appl Genet 94: 347–355.
  39. 39. Stace CA (2000) Cytology and cytogenetics as a fundamental taxonomic resource for the 20th and 21st centuries. Taxon: 451–477.
  40. 40. Martínez J. PV , Luceño M., and Cuadrado A. (2010) Evolution of Iris subgenus Xiphium based on chromosome numbers, FISH of nrDNA (5S, 45S) and trnL-trnF sequence analysis. Plant Syst Evol 289: 223–235.
  41. 41. Mouterde P (1983) Nouvelle flore du Liban et de la Syrie. Beyrouth (Liban): Imprimerie catholique.
  42. 42. The Plant List (2013) Version 1.1.: Published on the Internet;
  43. 43. Levan A, Fredga K, Sandberg AA (1964) Nomenclature for centromeric position on chromosomes. Hereditas 52: 201–220.
  44. 44. Peruzzi L, Eroglu H (2013) Karyotype asymmetry: again, how to measure and what to measure? Comparative cytogenetics 7: 1. pmid:24260685
  45. 45. Watanabe K, Yahara T, Denda T, Kosuge K (1999) Chromosomal evolution in the genus Brachyscome (Asteraceae, Astereae): statistical tests regarding correlation between changes in karyotype and habit using phylogenetic information. Journal of Plant Research 112: 145–161.
  46. 46. Paszko B (2006) A critical review and a new proposal of karyotype asymmetry indices. Plant Systematics and Evolution 258: 39–48.
  47. 47. Zarco CR (1986) A new method for estimating karyotype asymmetry. Taxon: 526–530.
  48. 48. Geber G, Schweizer D (1988) Cytochemical heterochromatin differentiation in Sinapis alba (Cruciferae) using a simple air-drying technique for producing chromosome spreads. Plant Syst Evol 158: 97–106.
  49. 49. Schweizer D (1976) Reverse fluorescent chromosome banding with chromomycin and DAPI. Chromosoma 58: 307–324. pmid:137107
  50. 50. Siljak-Yakovlev S, Cerbah M, Coulaud J, Stoian V, Brown SC, Zoldos V, et al. (2002) Nuclear DNA content, base composition, heterochromatin and rDNA in Picea omorika and Picea abies. Theor Appl Genet 104: 505–512. pmid:12582725
  51. 51. Gerlach WL, Dyer TA (1980) Sequence organization of the repeating units in the nucleus of wheat which contain 5S rRNA genes. Nucleic Acids Res 8: 4851–4865. pmid:7443527
  52. 52. Heslop-Harrison J (1991) In situ hybridization with automated chromosome denaturation. Technique 3: 109–116.
  53. 53. Peruzzi L, Eroğlu H (2013) Karyotype asymmetry: again, how to measure and what to measure? Comparative Cytogenetics 7.
  54. 54. Paszko B (2006) A critical review and a new proposal of karyotype asymmetry indices. Plant Syst Evol 258: 39–48.
  55. 55. Mitra J (1956) Karyotype Analysis of Bearded Iris. Botanical Gazette 117: 265–293.
  56. 56. Koçyiğit M, Erken K, Özhatay FN, Kaya E (2013) Iris L. Subgen. Iris Ve Subgen. Scorpiris Tabanında Kültür Bitkileri Üzerindeki Karyolojik Çalışmaların Önemi. V Süs Bitkileri kongresi. Yalova, Turkey. pp.
  57. 57. Wilson CA (2004) Phylogeny of Iris based on chloroplast matK gene and trnK intron sequence data. Mol Phylogen Evol 33: 402–412.
  58. 58. Hall T, Seisums A (2014) 793. Iris wallisiae. Curtis's Botanical Magazine 31: 238–248.
  59. 59. Tohmé G, Tohmé H (2002) A thousand and one flowers of Lebanon: Lebanese University.
  60. 60. Tohmé G, Tohmé H (2007) Illustrated flora of Lebanon. Beirut: The Lebanese University Publications Department. 309 p.
  61. 61. Tohmé G, Tohmé H (2011) Nouvelles recherches sur la flore endémique et naturalisée du Liban. Lebanese Science Journal 12: 133–141.
  62. 62. Simonet M (1932) Recherches cytologiques et génétiques sur les Iris. Bull Biol France Belg 78,: 696–707.
  63. 63. Simonet M (1934) Nouvelles recherches cytologiques et génétiques chez les iris. Ann sci nat, Bot Annales des sciences naturelles Botanique 25.
  64. 64. Siljak-Yakovlev S, Bogunić, F., Muratović, E., Šolić, M.E., Pavlović, D. & Medjedović, S. (2005) Genome organization of some Iris species assessed by molecular cytogenetics. XVII International Botanical Congress. Vienne. pp. 384.
  65. 65. Davis AP, Jury SL (1990) A taxonomic review of Iris L. series Unguiculares (Diels) Lawrence. Bot J Linn Soc 103: 281–300.
  66. 66. Goldblatt P, Takei M (1997) Chromosome cytology of Iridaceae-Patterns of variation, determination of ancestral base numbers, and modes of karyotype change. Annals of the Missouri Botanical Garden: 285–304.
  67. 67. Cerbah M, Souza-Chies T, Jubier M, Lejeune B, Siljak-Yakovlev S (1998) Molecular phylogeny of the genus Hypochaeris using internal transcribed spacers of nuclear rDNA: inference for chromosomal evolution. Mol Biol Evol 15: 345–354. pmid:9501501
  68. 68. Hidalgo O, Garcia-Jacas N, Garnatje T, Susanna A, Siljak-Yakovlev S (2007) Karyological evolution in Rhaponticum Vaill.(Asteraceae, Cardueae) and related genera. Bot J Linn Soc 153: 193–201.
  69. 69. Martel E, Poncet V, Lamy F, Siljak-Yakovlev S, Lejeune B, Sarr A (2004) Chromosome evolution of Pennisetum species (Poaceae): implications of ITS phylogeny. Plant Syst Evol 249: 139–149.
  70. 70. Siljak-Yakovlev S, Benmalek S, Cerbah M, DelaPeña TC, Bounaga N, Coba de la Pena T, et al. (1996) Chromosomal sex determination and heterochromatin structure in date palm. Sexual Plant Reproduction 9: 127–132.
  71. 71. Clarkson J, Lim K, Kovařík A, Chase M, Knapp S, Leitch A (2005) Long-term genome diploidization in allopolyploid Nicotiana section Repandae (Solanaceae). New Phytol 168: 241–252. pmid:16159337
  72. 72. Schubert I, Wobus U (1985) In situ hybridization confirms jumping nucleolus organizing regions in Allium. Chromosoma 92: 143–148.
  73. 73. Garcia S, Crhák Khaitová L, Kovařík A (2012) Expression of 5 S rRNA genes linked to 35 S rDNA in plants, their epigenetic modification and regulatory element divergence. BMC Plant Biol 12:95. pmid:22716941
  74. 74. Lacadena J, Cermeno M, Orellana J, Santos J (1984) Evidence for wheat-rye nucleolar competition (amphyplasty) in Triticale by silverstaining procedure. Theor Appl Genet 67: 207–213. pmid:24258550
  75. 75. Pikaard C (2001) Genomic change and gene silencing in polyploids. Trends Genet 17: 675–677. pmid:11718903
  76. 76. Lim K, Matyasek R, Kovarik A, Leitch A (2007) Parental Origin and Genome Evolution in the Allopolyploid Iris versicolor. Ann Bot 100: 219–224. pmid:17591610
  77. 77. Alvarez I, Wendel J (2003) Ribosomal ITS sequences and plant phylogenetic inference. Mol Phylogen Evol 29: 417–434.
  78. 78. Dadejova M, Lim K, Souckova´-Skalicka´ K, Matyasek R, Grandbastien M, Leith A, et al. (2007) Transciption activity of rRNA genes correlates with a tendency towards intergenomic homogenisation in Nicotiana allotetraploids. New Phytol 174: 658–668. pmid:17447920
  79. 79. Kovarik A, Pires J, Leitch A, Lim K, Sherwood A, Matyasek R, et al. (2005) Rapid concerted evolution of nuclear ribosomal DNAin two allopolyploids of recent and recurrent origin. Genetics 169: 931–944. pmid:15654116
  80. 80. Weiss-Schneeweiss H, Schneeweiss G, Stuessy T, Mabuchi T, Park J, Jang CG et al. (2007) Chromosomal stasis in diploids contrasts with genome restructuring in auto- and allopolyploid taxa of Hepatica (Ranunculaceae). New Phytol 174: 669–682. pmid:17447921