Ploidy Levels, Relative Genome Sizes, and Base Pair Composition in Cotoneaster

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Joseph J. Rothleutner Lincoln Park Zoo, 2001 North Clark Street, Chicago, IL 60614

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Mara W. Friddle Department of Horticulture, Oregon State University, 4017 Agricultural and Life Sciences Building, Corvallis, OR 97331

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Ryan N. Contreras Department of Horticulture, Oregon State University, 4017 Agricultural and Life Sciences Building, Corvallis, OR 97331

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Abstract

The genus Cotoneaster (Rosaceae, Maloideae) is highly diverse, containing ≈400 species. Like other maloids, there is a high frequency of naturally occurring polyploids within the genus, with most species being tetraploid or triploid. Apomixis is also prevalent and is associated with polyploidy. The objective of this study was to estimate genome sizes and infer ploidy levels for species that had not previously been investigated as well as compare estimates using two fluorochromes and determine base pair (bp) composition. Chromosome counts of seven species confirmed ploidy levels estimated from flow cytometric analysis of nuclei stained with 4′,6-diamidino-2-phenylindole (DAPI). Monoploid (1Cx) genome sizes ranged from 0.71 to 0.96 pg. Differences in monoploid genome size were not related to current taxonomic treatment, indicating that while chromosome sizes may vary among species, there are no clear differences related to subgeneric groups. A comparison of DAPI and propidium iodide (PI) showed a difference in DNA staining in Cotoneaster comparable to other rosaceous species. Base pair composition (AT%) in Cotoneaster ranged from 58.4% to 60.8%, which led to overestimation of genome size estimates in many cases—assuming the estimates of the DNA intercalator are accurate. Our findings will inform breeders with regard to the reproductive behavior of potential parents and may be used to confirm hybrids from interploid crosses.

Cotoneaster is a genus of woody plants composed of ≈400 species that range in habit from tight, impenetrable groundcovers to airy shrubs and medium-sized trees. The center of species diversity is the Himalayas and mountains of Yunnan and Sichuan provinces of China. The distribution encompasses the temperate zones of Eurasia and Northern Africa. The northern end of the range stretches from Spain to Siberia, and the southern limit extends from Morocco to the southern tip of India and South Korea (Fryer and Hylmö, 2009).

Although there are hundreds of species of Cotoneaster, a relatively small percentage are commonly grown in ornamental landscapes, as illustrated by Dirr (2009) listing only 14. These species were selected for their multiseason interest from flowers, fruit, and plant habit. In the 2014 Census of Horticultural Specialties (U.S. Department of Agriculture, 2014), Cotoneaster sales were estimated to exceed $7 million in the United States, although the value is likely greater because this figure accounted only for sales of Cotoneasters classified as “broadleaf evergreens” and many species are deciduous or semievergreen depending on climate and environmental factors.

Cotoneaster is a member of Rosaceae, subfamily Maloideae, and appears to be most closely related to Pyracantha (firethorn) and Heteromeles (christmas berry) (Robertson et al., 1991; Rohrer et al., 1992). Taxonomy at the family level is complicated, with interspecific and intergeneric hybridization being common. Interspecific hybrids of several species of Cotoneaster have been reported, and Cotoneaster melanocarpus has reportedly hybridized with Sorbus acuparia ssp. siberica to form the intergeneric hybrid ×Sorbocotoneaster (Fryer and Hylmö, 2009). Within Cotoneaster, there are two subgenera, Chaenopetalum and Cotoneaster, which are primarily defined by floral morphology. These subgenera have been further divided into 11 sections based on botanical characteristics, and further dissected into 37 series based on botanical characteristics and geographic origins of the species (Flinck and Hylmö, 1966). However, keys associated with this treatment are ambiguous and often of limited use for species identification. We are collaborating with Hoyt Arboretum (Portland, OR) to identify and evaluate our germplasm collection, with little success in identifying unknown samples.

The base chromosome number of Maloideae is 17 and is thought to be of allopolyploid origin—perhaps derived from a hybridization event between other subfamilies in Rosaceae [Rosoideae (x = 7, 8, 9), Spiraeoideae (x = 9), Amygaloideae (x = 8)] followed by a whole genome doubling event (Dickson et al., 1992; Sax, 1954). Cotoneaster species show a ploidy series, with estimates of 70% tetraploid (2n = 4x = 68), 15% triploid (2n = 3x = 51), and 10% diploid (2n = 2x = 34), and the remaining species of greater ploidy level (Fryer and Hylmö, 2009). Apomixis is common in Cotoneaster and appears to be associated with polyploidy, as the tetraploids and triploids are frequently obligate or rarely facultative apomicts, while diploid progeny are sexually derived (Bartish et al., 2001; Czapik, 1996; Hjelmqvist, 1962; Nybom and Bartish, 2007).

Because apomixis is so common in polyploid Cotoneaster, knowledge of ploidy level is essential for breeders to design crosses with hopes of hybrid seed, as the female must be a sexually fertile diploid. In addition, information on ploidy level, genome size, and bp composition may give taxonomists and phylogeneticists insight to the evolution and organization of the genus and related taxa. Previous reports of genome sizes in Cotoneaster are limited; therefore, our goals were to determine relative genome sizes and produce ploidy estimates across a wide selection of Cotoneaster including its breadth of taxonomic groups.

Materials and Methods

Plant material.

Germplasm was collected through various means including whole plants from nurseries, cuttings from gardens and arboreta, and seeds from gardens around the world participating in Index Seminum (Table 1). The latter formed the bulk of our collection. Plants were maintained in containers or in field plots at Oregon State University and all were assigned accession numbers.

Table 1.

Source and collection information for 67 Cotoneaster accessions.

Table 1.

Genome sizing.

Holoploid (2C) genome sizes were determined by flow cytometry (CyFlow PA; Partec, Münster, Germany) and comparison of mean relative fluorescence of the sample against an internal standard, Pisum sativum ‘Ctirad’, with a known genome size of 8.76 pg (Greilhuber et al., 2007). Two different fluorochromes were used. A total of 67 accessions representing 65 species were sampled across the two subgenera and all 11 sections using flow cytometric analysis of nuclei stained with DAPI (CyStain ultraviolet Precise P; Partec). A subset of 17 taxa was also prepared with PI (CyStain ultraviolet Absolute P; Partec). Nuclei of each sample and our standard were concurrently prepared by chopping with a double-sided razor blade in extraction buffer (CyStain ultraviolet Precise P Nuclei Extraction Buffer; Partec) for ≈90 s before being filtered through a 50-μm nylon mesh filter (CellTrics®; Partec) and stained with either fluorochrome. For PI-stained samples, RNase was included to ensure staining of DNA exclusively. Cotoneaster samples were prepared using 4 cm of rapidly growing terminal stem tissue including vegetative buds and we used 1 cm2 of fresh pea leaf tissue. DAPI-stained samples were incubated in darkness for 5 to 10 min before analysis and PI-stained samples were incubated in darkness for at least 30 min on ice. Three replicates of each accession were prepared for both DAPI and PI. A minimum of 3000 particles were analyzed for each sample. Sample runs were rejected if the coefficient of variation (cv) was greater than 7%.

Holoploid DNA content (2C) was calculated as DNA content of standard × (mean fluorescence value of sample/mean fluorescence of standard). Then, analysis of variance and means separation by Tukey’s honestly significant difference was performed, with ploidy levels inferred from mean separation. Monoploid genome sizes were calculated by dividing each sample’s 2C genome size by inferred ploidy. Analysis of variance was then conducted on monoploid genome size by accession and then taxonomic division to test for significant differences among subgenera or sections; mean separation was performed as described above when the model was significant (α = 0.05). For the subset of 17 accessions that were examined with both DAPI and PI, genome size estimates using each fluorochrome were compared using a t test (α = 0.05) separately for each accession. Base pair composition was calculated as AT% = AT% for internal standard × {[(fluorescence sample, DAPI)/(fluorescence internal standard, PI/fluorescence sample, PI)](1/binding length)} (Godelle et al., 1993). AT% of P. sativum ‘Ctirad’ is 61.50% and has a binding length of ≈3.5 bp (Meister and Barrow, 2007).

Cytology.

Chromosomes were counted for seven species, five of which were included in the genome sizing, and two additional species Cotoneaster hebephyllus and Cotoneaster poluninii. Somatic cells were collected from actively growing root tips, which grew freely from the bottom of their containers into sand. Roots were treated with 0.003 m 8-hydroxyquinoline for 2 h at 4 °C and fixed in Carnoy’s solution [6 absolute ethanol : 3 chloroform : 1 glacial acetic acid (by volume)] overnight. Root tips were stored in 70% ethanol at 4 °C until prepared for chromosome counts (Goldblatt and Gentry, 1979). Chromosomes were examined by root tip squashes with modified carbol fuchsin, at ×63 to ×100 magnification (Axio imager.A1; Zeiss, Thornwood, NY) and images were collected using a monochromatic CCD camera (AxioCam MRm; Zeiss). A minimum of three cells were counted for each species.

Results and Discussion

Relative 2C genome sizes for 67 accessions determined by flow cytometry with DAPI ranged from 1.52 pg (Cotoneaster frigidus) to 4.71 pg (Cotoneaster kweitschoviensis) (Table 2). The 2C genome sizes showed marked divisions, which were used to assign ploidy level. Of the 67 accessions, 10 (15%) were diploids (2n = 2x = 34) with 2C values ranging from 1.52 to 1.73 pg, 5 (9%) were triploids (2n = 3x = 51) with 2C values ranging from 2.14 to 2.58 pg, 51 (76%) were tetraploids (2n = 4x = 68) with 2C values ranging from 2.88 to 3.34 pg, and 1 (1.5%) accession was hexaploid (2n = 6x = 102) with a 2C value of 4.71 pg (Table 2). Regarding relative percentage of ploidy levels, our findings generally agree with previous reports including Kroon (1975) who reported 3 diploid, 3 triploid, and 23 tetraploid species among the 28 studied. Ours is the first report of ploidy estimation for 13 of the species. However, one diploid species, Cotoneaster juratana, is not a valid species and we have been unable to confirm its identity. One assumption was a mislabeling of Cotoneaster juranus, which has only been reported as a tetraploid (Fryer and Hylmö, 2009). Unfortunately, morphology of the plant labeled as C. juratana did not match the description of C. juranus and has since been lost from our collection. We sampled two accessions of Cotoneaster adpressus and Cotoneaster acutifolius and both showed ploidy series. Cotoneaster adpressus 10-0157 was tetraploid and the other, ‘Tom Thumb’, was triploid. Previous reports for C. adpressus indicated that it was either diploid or triploid (Sax, 1954; Zeilinga, 1964), making ours the first report of tetraploidy for the species. Cotoneaseter acutifolius 09–0047 was diploid and C. acutifolius 10–0126 was triploid. Our results are consistent with Sax (1954) who reported diploids, triploids, and tetraploids for C. acutifolius. This is in contrast to Zeilinga (1964) who reported that all species in their study were diploid or tetraploid and only cultivars were found to be triploid. This provides evidence that ‘Tom Thumb’ and C. acutifolius 10-0126 are hybrids. Cotoneaster ×watereri 10-106 was a triploid in contrast to previous reports indicating this hybrid species to be diploid (Fryer and Hylmö, 2009). This species arose as a hybrid of C. frigidus × Cotoneaster salicifolius, which generally are both regarded as diploids, though Sax (1954) also reported triploidy in C. salicifolius. It is unclear how this triploid accession arose, but possibilities include a previously unknown tetraploid cytotype of one of the parent species, unreduced gamete production in one of the progenitors, an apomictic seedling, or even a self-pollinated seedling from an unreported triploid cytotype of C. salicifolius, as triploids often are regarded as facultative apomicts. Another alternative is that the parent plant from which we received seed was diploid and it was accidentally pollinated by a tetraploid, resulting in triploid progeny. There is some evidence against obligate apomixis in C. ×watereri, as Fryer and Hylmö (2009) describe five cultivars, a level of diversity that would not be expected if the species was an obligate apomict. Both Zeilinga (1964) and Kroon (1975) reported that all species were diploid or tetraploid and triploids only arise through hybridization. To determine if that assertion is supported by our study, a detailed morphological or molecular investigation of species and hybrids would be required, which is beyond the scope of the current research. In five other species, our data indicated a different cytotype than reported by Fryer and Hylmö (2009). These include Cotoneaster ganghobaensis, C. kweitschoviensis, Cotoneaster milkedandaensis, Cotoneaster niger, and Cotoneaster sikagensis. The differing cytotypes we report do not suggest consistent error in either direction (over or underestimates) and include two 4x that were reportedly 3x, one 6x previously reported as 4x, one 4x previously reported as 2x, and one 2x previously reported as 4x (Table 2). A larger screening with more accessions representing each species would likely uncover more ploidy series within other species as well as identifying more examples for which new findings would differ from previous reports. Discrepancies in ploidy between previous papers and our findings are due to testing different sources or accessions of material, some of which may have arisen from hybridization.

Table 2.

Holoploid (2C) and monoploid (1Cx) genome sizes of 67 accessions of Cotoneaster determined by flow cytometric analysis of nuclei stained with 4′,6-diamidino-2-phenylindole (DAPI) using Pisum sativum ‘Ctirad’ (2C = 8.76 pg) as an internal standard, inferred ploidy levels based on flow cytometry data, previously reported ploidy, and mean 1Cx value of 10 taxonomic sections studied.

Table 2.

When looking across taxonomic divisions, variation in ploidy level was observed in the subgenera and within many sections (Table 2). The 1Cx genome size ranged from 0.71 pg of DNA for C. acutifolius (10-0126) to 0.87 pg of DNA for Cotoneaster dammeri. When compared across all taxa, monoploid genome sizes showed a detectable difference (P < 0.05); however, when these differences were examined by taxonomic division, they were insignificant (P ≥ 0.05). Since we did not observe differences between taxonomic groups, we infer that our mean monoploid genome size can be used to calibrate our ploidy estimations in future investigations. In addition, when compared with other genera like Magnolia, which has observable differences in monoploid genome size between taxonomic sections (Parris et al., 2010), Cotoneaster has undergone relatively little divergence in chromosome size.

DAPI genome size estimates generally were larger than PI with differences ranging from 0.03 to 0.14 pg (Table 3). Fifteen of 17 accessions were different (P < 0.05) using respective fluorochromes and four accessions had higher significance (P < 0.0001). Doležel et al. (1992) reported significant differences (P < 0.01) for five genera in four families including four genera for which DAPI overestimated genome size from 11% to 30% compared with PI and one species for which DAPI underestimated genome size by nearly 27%. Our calculations show that bp composition in Cotoneaster ranges from 58.4% to 60.8% AT (Table 3), which agrees with estimates of 59.2% to 61.1% AT for three genera from Rosaceae (Meister and Barrow, 2002). We found no relationship between ploidy level (thus genome size) and AT%, a finding consistent with previous reports in Rosaceae (Meister and Barrow, 2002). Interestingly, we observed a trend that the lower the AT%, the greater the overestimation of genome size using DAPI compared with PI. Even though Parris et al. (2010) found that DAPI underestimated genome size for Magnolia, both of our studies show the same trend: increasing AT% results in a lower DAPI estimate compared with PI. Although the trends were similar, we observed overestimation using DAPI, whereas Parris et al. (2010) observed underestimation, even though AT% was similar, albeit higher in their study. A possible source of variation in our study was using different types of tissue in our samples (young stems and vegetative buds) and pea standard (young, expanded leaves). It is possible this resulted in variation in chromatin structure, which would affect the amount of unstainable DNA (Doležel et al., 1992).

Table 3.

Monoploid (1Cx) genome sizes determined by analysis of nuclei stained with 4′,6-diamidino-2-phenylindole (DAPI) or propidium iodide (PI) using Pisum sativum ‘Ctirad’ (2C = 8.76 pg) as an internal standard, the difference between genome size estimates between fluorochromes and bp composition of 17 taxa of Cotoneaster.

Table 3.

Although our choice of fluorochrome influenced genome size estimates, most differences were not large enough to affect ploidy estimation. However, there could be some confusion for several species included in our analysis. For instance, Cotoneaster boisianus showed a 0.14 pg difference in monoploid genome size estimate between fluorochromes. Using the estimate from PI of 2.72, the inferred ploidy level would calculate as 3.5x. Without additional information provided from cytological analysis, accurate ploidy assignment may be challenging and sample readings could erroneously be interpreted as aneuploid. DAPI is less expensive, uses less toxic compounds, and often resulted in lower cv for mean nuclei fluorescence than PI for Cotoneaster. Overall, we consider DAPI acceptable for our purposes in an applied breeding program.

From our chromosome counts in seven species, we found one diploid accession and six tetraploid accessions (4x Cotoneaster vandelarii not shown; Fig. 1). These ploidy estimates matched our results from flow cytometry for the five species that were examined by both methods. Overall, when our results were compared with literature for both cytology and flow cytometry, most were in agreement. Where there is conflict in the reports, the conflict may be from the way that the ploidy reports have been generated. Sax (1954) conducted estimations via chromosome counting in pollen mother cells and Zeilinga (1964) found several conflicting reports when root tips were examined. Zeilinga suggested that with polyploids, the chromosome pairing observed in pollen mother cells was crowded and led to confusion in counting. With a relatively high base chromosome number and common occurrence of polyploidy (102 chromosomes in hexaploids), it is possible that previous chromosome counts included errors. Determining ploidy level by counting chromosomes in Cotoneaster was time consuming and difficult, while flow cytometry proved to be much faster and accurate.

Fig. 1.
Fig. 1.

Photomicrographs of six Cotoneaster accessions including (A) a diploid (2n = 2x = 34) Cotoneaster henryanus, and (B) five tetraploids (2n = 4x = 68) Cotoneaster buxifolius, (C) Cotoneaster vandelarii, (D) Cotoneaster hebephyllus, (E) Cotoneaster lidjiangensis, and (F) Cotoneaster poluninii. Cells were prepared from root tips and chromosomes were stained using modified carbol fuchsin.

Citation: Journal of the American Society for Horticultural Science J. Amer. Soc. Hort. Sci. 141, 5; 10.21273/JASHS03776-16

Thus far, taxonomic organization in Cotoneaster has largely failed to incorporate molecular data and has relied on morphology and species provenance, although a report by Bartish et al. (2001) using randomly amplified polymorphic DNA supported the recognition of subgenera. Because of the number of species within the genus and the difficulty in organizing Cotoneaster, we hope fundamental information on genome size, ploidy level, and bp composition may give others insight to the relationship among the species. However, the results of this study do not show a relationship of chromosome size to current taxonomic organization, as monoploid genome sizes did not appear to be linked to taxonomic division.

This work may be useful to breeders for predicting success of interspecific hybridization and fertility of F1 populations. Along with other factors, similarity in chromosome size contributes to functional meiosis and bivalent pairing between genomes. In this study, monoploid genome sizes varied up to 23% among species. In Rudbeckia, a hybrid was recovered when there was a difference in genome size of >300% (Palmer et al., 2009). The much smaller range in Cotoneaster suggests variation in monoploid genome size, thus chromosome size, is not expected to hinder interspecific hybridization. Furthermore, we have successfully performed several intersubgeneric and interploidy crosses including Cotoneaster ×suecicus ‘Coral Beauty’ (2x, 2C = 1.53 pg, subgenus Chaenopetalum) × Cotoneaster splendens (4x, 2C = 3.03 pg, subgenus Cotoneaster) that resulted in a triploid hybrid, which was confirmed using flow cytometry (2C = 2.42 pg), thus supporting broad compatibility in the genus. Breeding programs should conduct ploidy analysis for each accession included in a germplasm collection without relying on previous reports. We expect the broader the survey among species from various sources, the more examples of ploidy series will be found. Nevertheless, we have demonstrated the utility of apomictic polyploids for breeding when used as pollen parents.

Taxonomy of Cotoneaster is challenging due to morphological similarities, the propensity for hybridization, and the presence of apomixis. The tangled issue of separating and correctly identifying these species is emphasized by Dirr (2009) who stated, “Cotoneaster identification is not easy with 400 species, many possibly the result of hybridization, subsequent apomixis, which leads to microspecies that essentially reproduce vegetatively via seed.” Our study relied heavily on material obtained through Index Seminum and here we present the material labeled as we received it. Because of the presence of apomixis, particularly among tetraploids, we have strong confidence in the identification of most species presented. It is worth noting that materials in our study were not wild collected as one may expect in a classical floristic study but our findings should be quite relevant to applied plant breeders or others studying cultivated material of Cotoneaster or Maloideae.

Literature Cited

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  • Bartish, I.V., Hylmö, B. & Nybom, H. 2001 RAPD analysis of interspecific relationships in presumably apomictic Cotoneaster species Euphytica 120 273 280

    • Search Google Scholar
    • Export Citation
  • Czapik, R. 1996 Problems of apomictic reproduction in the families Compositae and Rosaceae Folia Geobot. Phytotaxon. 31 381 387

  • Darlington, C.D. & Wylie, A.P. 1956 Chromosome atlas of flowering plants. Macmillan, New York, NY

  • Dickson, E.E., Arumunganathan, K., Kresovich, S. & Doyle, J.J. 1992 Nuclear DNA content variation within the Rosaceae Amer. J. Bot. 79 1081 1086

  • Dirr, M.A. 2009 Manual of woody landscape plants. 6th ed. Stipes Publ., Champaign, IL

  • Doležel, J., Sgorbati, S. & Lucretti, S. 1992 Comparison of three DNA fluorochromes for flow cytometric estimation of nuclear DNA contents in plants Physiol. Plant. 85 625 631

    • Search Google Scholar
    • Export Citation
  • Flinck, K.E. & Hylmö, B. 1966 A list of series and species in the genus Cotoneaster Bot. Not. 119 445 463

  • Fryer, J. & Hylmö, B. 2009 Cotoneasters: A comprehensive guide to shrubs for flowers, fruit and foliage. Timber Press, Portland, OR

  • Godelle, B., Cartier, D., Marie, D., Brown, S.C. & Siljak-Yakovlev, S. 1993 Heterochromatin study demonstrating the non-linearity of fluorometry useful for calculating genomic base composition Cytometry 14 618 626

    • Search Google Scholar
    • Export Citation
  • Goldblatt, P. & Gentry, A.H. 1979 Cytology of Bignoniaceae Bot. Not. 132 475 482

  • Greilhuber, J., Temsch, E.M. & Loureiro, J.C.M. 2007 Nuclear DNA content measurement, p. 67–101. In: J. Doležel, J. Greilhuber, and J. Suda (eds.). Flow cytometry with plant cells: Analysis of genes, chromosomes and genomes. Wiley-VCH, Weinheim, Germany

  • Hjelmqvist, H. 1962 Embryo sac development of Cotoneaster Bot. Not. 14 209 236

  • Jedrzejczyk, I. & Sliwinska, E. 2010 Leaves and seeds as materials for flow cytometric estimation of the genome size of 11 Rosaceae woody species containing DNA-staining inhibitors J. Bot. 2010 1 9

    • Search Google Scholar
    • Export Citation
  • Kroon, G.H. 1975 Polyploidy in Cotoneaster II Acta Botanisker Neerlandica 24 417 420

  • Meister, A. & Barrow, M. 2002 Lack of correlation between AT frequency and genome size in higher plants and the effect of nonrandomness of base sequences on dye binding Cytometry 47 1 7

    • Search Google Scholar
    • Export Citation
  • Meister, A. & Barrow, M. 2007 Analysis of genes, chromosomes and genomes, DNA base composition of plant genomes, p. 177–185. In: J. Doležel, J. Greilhuber, and J. Suda (eds.). Flow cytometry with plant cells: Analysis of genes, chromosomes and genomes. Wiley-VCH, Weinheim, Germany

  • Nybom, H. & Bartish, I.V. 2007 DNA markers and morphometry reveal multiclonal and poorly defined taxa in an apomictic Cotoneaster species complex Taxon 56 119 128

    • Search Google Scholar
    • Export Citation
  • Palmer, I.E., Ranney, T.G., Lynch, N.P. & Bir, R.E. 2009 Crossability, cytogenetics and reproductive pathways in Rudbeckia subgenus Rudbeckia HortScience 44 44 48

    • Search Google Scholar
    • Export Citation
  • Parris, K.A., Ranney, T.G., Knap, H.T. & Baird, W.V. 2010 Ploidy levels, relative genome sizes, and base pair composition in magnolia J. Amer. Soc. Hort. Sci. 135 533 547

    • Search Google Scholar
    • Export Citation
  • Robertson, K.R., Rohrer, J.R., Phipps, J.B. & Smith, P.G. 1991 A synopsis of genera in the subfamily Maloideae (Rosaceae) Syst. Bot. 16 376 394

  • Rohrer, J.R., Robertson, K.R. & Phipps, J.B. 1992 Variation in structure among fruits of Maloideae (Rosaceae) Amer. J. Bot. 78 1617 1635

  • Sax, H.J. 1954 Polyploidy and apomixis in Cotoneaster J. Arnold Arbor. 35 334 365

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Joseph J. Rothleutner Lincoln Park Zoo, 2001 North Clark Street, Chicago, IL 60614

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Mara W. Friddle Department of Horticulture, Oregon State University, 4017 Agricultural and Life Sciences Building, Corvallis, OR 97331

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Ryan N. Contreras Department of Horticulture, Oregon State University, 4017 Agricultural and Life Sciences Building, Corvallis, OR 97331

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Contributor Notes

This study is from a thesis submitted by Joseph. J. Rothleutner as partial fulfillment of the degree of master of science in Horticulture at Oregon State University.

This research was funded, in part, by the Oregon Agricultural Experiment Station and by USDA-ARS Specific Cooperative Agreement 58-1230-3-501.

We are grateful for germination and early maintenance of plants by Imogene Hollis and Heather Stoven. Also, we extend our gratitude to contributing gardens and nurseries for generously supplying plants or seeds.

Corresponding author. E-mail: ryan.contreras@oregonstate.edu.

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  • Photomicrographs of six Cotoneaster accessions including (A) a diploid (2n = 2x = 34) Cotoneaster henryanus, and (B) five tetraploids (2n = 4x = 68) Cotoneaster buxifolius, (C) Cotoneaster vandelarii, (D) Cotoneaster hebephyllus, (E) Cotoneaster lidjiangensis, and (F) Cotoneaster poluninii. Cells were prepared from root tips and chromosomes were stained using modified carbol fuchsin.

 

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