# Evolutionary insights into the genomic organization of major ribosomal DNA in ant chromosomes

Gisele Amaro Teixeira, Hilton Jeferson Alves Cardoso de Aguiar, Frédéric Petitclerc, Jérôme Orivel, Denilce Meneses Lopes, Luisa Antonia Campos Barros

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## To cite this version:

Gisele Amaro Teixeira, Hilton Jeferson Alves Cardoso de Aguiar, Frédéric Petitclerc, Jérôme Orivel, Denilce Meneses Lopes, et al.. Evolutionary insights into the genomic organization of major ribosomal DNA in ant chromosomes. Insect Molecular Biology, 2021, 30 (3), pp.340-354. 10.1111/imb.12699. hal-03369943

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Submitted on 7 Oct 2021

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Distributed under a Creative Commons Attribution 4.0 International License
1 Evolutionary insights into the genomic organization of major ribosomal

2 DNA in ant chromosomes

3

4 Gisele Amaro Teixeira¹, ², Hilton Jeferson Alves Cardoso de Aguiar³*, Frédéric

5 Petitclerc⁴, Jérôme Orivel⁴, Denilce Meneses Lopes², Luísa Antônia Campos Barros³

6

7 1 Programa de Pós-graduação em Biologia Celular e Estrutural, Universidade Federal

8 de Viçosa, Viçosa, MG 36570-000, Brazil.

9 2 Laboratório de Citogenética de Insetos, Departamento de Biologia Geral,

10 Universidade Federal de Viçosa, Viçosa, MG 36570-900, Brazil.

11 3-Universidade Federal do Amapá, Campus Binacional – Oiapoque, n°3051, Bairro

12 Universidade, 68980-000, Oiapoque, Amapá, Brazil.

13 4 CNRS, UMR EcoFoG, AgroParisTech, CIRAD, INRA, Université de Guyane,

14 Université des Antilles, Campus Agronomique, BP 316, 97379 Kourou Cedex, France.

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16 Short running title: Evolution of rDNA genes in ant chromosomes

17 * Corresponding author

18 E-mail: hilton.aguiar@hotmail.com
1 Abstract

2 The major rDNA genes are composed of tandem repeats and are part of the nucleolus
3 organizing regions (NORs). They are highly conserved and therefore useful in
4 understanding the evolutionary patterns of chromosomal locations. The evolutionary
5 dynamics of the karyotype may affect the organization of rDNA genes within
6 chromosomes. In this study, we physically mapped 18S rDNA genes in 13 Neotropical
7 ant species from four subfamilies using fluorescence *in situ* hybridization. Furthermore,
8 a survey of published rDNA cytogenetic data for 50 additional species was performed,
9 which allowed us to detect the evolutionary patterns of these genes in ant chromosomes.

10 Species from the Neotropical, Palearctic, and Australian regions, comprising a total of 63
11 species from 19 genera within six subfamilies, were analyzed. Most of the species (48 out
12 of 63) had rDNA genes restricted to a single chromosome pair in their intrachromosomal
13 regions. The position of rDNA genes within the chromosomes appears to hinder their
14 dispersal throughout the genome, as translocations and ectopic recombination are
15 uncommon in intrachromosomal regions because they can generate meiotic
16 abnormalities. Therefore, rDNA genes restricted to a single chromosome pair seem to be
17 a plesiomorphic feature in ants, while multiple rDNA sites, observed in distinct
18 subfamilies, may have independent origins in different genera.

19

20 **Keywords:** Evolution - ribosomal DNA - gene dispersion - Formicidae - chromosome
21 rearrangements - physical mapping
# 1 Introduction

Eukaryotic genomes have repetitive tandem sequences such as in the major ribosomal RNA genes (45S = 18S + 5.8S + 28S), herein denominated rDNA, which contain highly conserved genic sequences and are therefore useful as molecular genetic markers, allowing comparisons across distant taxa. However, intergenic spacers vary in both sequence and length (Long and Dawid, 1980; Sumner, 2003; Symonová, 2019). The 45S ribosomal genes are part of the nucleolus organizing regions (NORs) and are located in portions of the DNA that, after their condensation, usually appear as secondary constrictions on metaphase chromosomes (Sumner, 2003).

The silver nitrate impregnation technique (Ag-NOR banding) shows only transcriptionally active NORs by staining nuclear acid proteins involved in transcription (Howell and Black, 1980; reviewed by Sumner, 2003). Species with multiple rDNA clusters do not usually exhibit silver-staining in all clusters (reviewed by Sumner, 2003; Vicari *et al.*, 2008). However, it must be assumed that if a species has only a single NOR (or single 45S rDNA gene site), then it will be transcriptionally active, as evidenced for different organisms (Dobigny *et al.*, 2002; Barros *et al.*, 2015; Falcione *et al.*, 2018; Cholak *et al.*, 2020; Malimpensa *et al.*, 2020).

Intercellular and interindividual variations are frequently detected by the Ag-NOR banding method (Zurita *et al.*, 1997; Cross *et al.*, 2003; Walker *et al.*, 2014; Schmid *et al.*, 2017). However, the use of this technique in ant species has not produced reliable results owing to low repeatability, difficulty in obtaining good-quality markings, and the appearance of unspecified marks on heterochromatic regions of many chromosomes, which makes using this procedure unsatisfactory and inconclusive (Imai *et al.*, 1992; Hirai *et al.*, 1994; Lorite *et al.*, 1997; Barros *et al.*, 2009, 2015; Aguiar *et al.*, 2017).
Since the 1980s, molecular cytogenetic tools have been used to study karyotypes.

For example, fluorescence *in situ* hybridization (FISH) has proven to be an effective and

precise tool for physically mapping specific DNA sequences within chromosomes

(reviewed by Levsky and Singer, 2003; Liehr, 2017). The rDNA genes can be located in

single or multiple chromosome pairs (Sochorová *et al.*, 2018). In several organisms,

studies of these genes have pointed to chromosomal differences within species complexes

(Mantovani *et al.*, 2005; Barbosa *et al.*, 2017; Dutra *et al.*, 2020) and between species

with similar karyotypes (Panzera *et al.*, 2012; Golub *et al.*, 2015; Gokhman *et al.*, 2016).

As a consequence, inferences can be made based on chromosomal rearrangements that

shape the chromosomal evolution of a species (Roy *et al.*, 2005; Nguyen *et al.*, 2010;

Britton-Davidian *et al.*, 2011; Cabral-de-Mello *et al.*, 2011; Dutrillaux and Dutrillaux,

2012; Roa and Guerra, 2012; Menezes *et al.*, 2019; Degrandi *et al.*, 2020).

In ants, the physical mapping of rDNA genes using the FISH technique was first

described in Australian ants by Hirai *et al.* (1994, 1996). Since then, the number of studies

mapping these genes has increased (Mariano *et al.*, 2008; Santos *et al.*, 2016; Micolino

*et al.*, 2019a; Teixeira *et al.*, 2020) and other repetitive sequences, such as telomeres

(Meyne *et al.*, 1995; Pereira *et al.*, 2018; Micolino *et al.*, 2020; Castro *et al.*, 2020),

satellite DNA (Lorite *et al.*, 2004; Huang *et al.*, 2016), 5S ribosomal genes (Aguiar *et al.,

2017), and microsatellites (Barros *et al.*, 2018; Micolino *et al.*, 2019b), have been mapped

in the chromosomes using the FISH technique. To date, molecular cytogenetic studies on

rDNA genes in ants have improved understanding of chromosomal evolution and

phylogeny, and provided taxonomic resolutions for different ant groups (Hirai *et al.,

1994, 1996; Santos *et al.*, 2010, 2016).

In this study, we physically mapped 18S rDNA clusters using the FISH technique

and verified if they were GC-rich in 13 Neotropical ant species from four subfamilies
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(Ectatomminae, Formicinae, Myrmicinae, and Ponerinae). In addition, we reviewed previous molecular cytogenetic data related to rDNA gene clusters (45S, 18S, or 28S) in ants. Using these data, we investigated whether the number and location of the ribosomal gene clusters followed a specific pattern or were randomly distributed in order to understand the genomic organization and evolutionary dynamics of these genes in ants.

# Results

## 1 - Chromosome mapping of 18S rDNA clusters in 13 Neotropical ant species

All of the studied ant species presented only a single 18S rDNA site that was co-localized with GC-rich regions ($CMA_3^+$) (Table 2), while AT-rich regions (DAPI$^+$) were not detected in any species. Most of the species presented GC-rich rDNA genes in the intrachromosomal regions (pericentromeric or interstitial), including *Pseudoponera gilberti* (Kempf, 1960) (Fig. 1A, S1A), *Anochetus targionii* Emery, 1894 (Fig. 1B, S1B), *Odontomachus haematodus* (Linnaeus, 1758) (Fig. 1C, D, S1C), *Odontomachus bauri* Emery, 1892 (Fig 1E, S1D), *Pheidole germaini* Emery, 1896 (Fig. 2A, S2A), *Crematogaster longispina* Emery, 1890 (Fig. 2B, S2B), *Solenopsis geminata* (Fabricius, 1804) (Fig. 2C, D, S2C), *Myrmicocrypta* sp. (Fig. 2E, S2D), and *Acromyrmex echinatior* (Forel, 1899) (Fig. 2F). Additional GC-rich bands only occurred in *P. gilberti* in the pericentromeric region of the 3rd and 4th metacentric chromosome pairs (Fig. S1B).
Differences in rDNA cluster sizes between homologous chromosomes were observed in heterozygous *O. bauri* individuals. One of the homologous chromosomes showed clusters approximately twice the size of those in the other chromosome (Fig. 1F, S1E). A male
1 individual bearing a chromosome with minor GC-rich bands was analyzed (Fig. S1F).

2 Homozygous individuals with duplicated clusters were not observed.

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<figure><img src="image_3.png" /><figcaption>**Figure 1.** Fluorescence *in situ* hybridization with 18S rDNA probe (red blocks) of Ponerinae ants: (A) *Pseudoponera gilberti* (2n=12), (B) *Anochetus targionii* (2n=30), (C, D) *Odontomachus haematodus* (2n=44, Brazil and French Guiana, respectively), (E, F) *Odontomachus bauri* (2n=44). In the last species, the 18S rDNA site was located on the long arm of the polymorphic chromosomal pair. The boxes show a polymorphic subtelocentric pair in (E) in the homozygous state, and (F) in the heterozygous state. Bars = 5 µm.</figcaption></figure>
<figure><img src="image_4.png" /><figcaption>**Figure 2.** Fluorescence *in situ* hybridization with 18S rDNA probe (red blocks) of Myrmicinae ants: (A) *Pheidole germaini* (2n=22), (B) *Crematogaster longispina* (2n=24), (C) *Solenopsis geminata* (2n=32) and (D) *Solenopsis geminata* (n=16), (E) *Myrmicocrypta* sp. (2n=30), (F) *Acromyrmex echinatior* (2n=38), (G) *Strumigenys diabola* (2n=40). Bars = 5 µm.</figcaption></figure>

The remaining species showed GC-rich 18S rDNA clusters across the entire chromosome arm, occupying either the long arm, as in *Gnamptogenys tortuolosa* (Smith, 1858) (Fig. 3A, S3A), or residing in the short arm, as in *Strumigenys diabola* Bolton, 2000 (Fig. 2G, S2E), *Camponotus atriceps* (Smith, 1858) (Fig. 3B, S3B), and *Gigantiops destructor* (Fabricius, 1804) (Fig. 3C, S3C, D). Heteromorphism of 18S rDNA clusters was detected in all of the analyzed *C. atriceps* and *G. tortuolosa* individuals. In the latter species, the heteromorphism of the NOR resulted in differences in total size between
1 homologous chromosomes, which changed their morphology such that one was
2 submetacentric while the other was subtelocentric. In *G. destructor*, additional GC-rich
3 bands were located in the interstitial region of the long arm of the largest subtelocentric
4 chromosome pair (Fig. S3C, D).
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<caption>**Figure 3.** Fluorescence *in situ* hybridization with 18S rDNA probe (red blocks) of an Ectatomminae ant (A) *Gnamptogenys tortuolosa* (2n=44), and Formicinae ants (B) *Camponotus atriceps* (2n=40) and (C) *Gigantiops destructor* (n=39). In *G. tortuolosa*, the box shows a remarkable size heteromorphism of 18S rDNA clusters between homologs. Bars: 5 µm.</caption>

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## 2 - Chromosome mapping review of rDNA clusters in ants

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17 Cytogenetic data available in the literature related to the rDNA genes of 50 ant species from 12 genera and six subfamilies were reviewed (Table 2; Fig. 4). Most data were concentrated on Neotropic ants, with information on 33 species, while the Palearctic
and Australian regions had data on only one and 16 species, respectively. A single rDNA site localized in the intrachromosomal region was observed in most species (Fig. 4A, B).

However, *Camponotus renggeri* Emery, 1894, *Dinoponera gigantea* (Perty, 1833), and most of the studied *Myrmecia* species presented multiple rDNA sites over the entire short chromosome arm. The subfamily Myrmicinae possessed most of the rDNA data, and the Myrmeciinae subfamily showed a different pattern in relation to other subfamilies with multiple rDNA sites observed in the majority of species (Fig. 4C). *Dolichoderus voraginosus* Mackay, 1993 did not show any co-localization of 18S rDNA clusters and GC-rich bands. Figure 5 summarizes the available data on the number and position of NORs in ant species and the phylogenetic relationships among these species, and is based on the published molecular phylogenies.

## Discussion

### 1 - Patterns of rDNA clusters in the karyotypes of specific ant groups

Specific rDNA patterns can be observed in the karyotypes of some ant groups where several species have been studied. In species of the genus *Dolichoderus*, the chromosome numbers range from 2n=10 to 58, and a single rDNA site in the interstitial region has been observed in the majority of the investigated species. However, there are two exceptions: *D. voraginosus* and *Dolichoderus attelaboides* (Fabricius, 1775), where the rDNA clusters are located in the terminal region of the long arm and in the short arm, respectively (Santos *et al.*, 2016).

Despite the chromosomal variation observed in *Gnamptogenys* spp. (2n=24 to 44), they all have a single rDNA site in the intrachromosomal region (Teixeira *et al.*, 2020).
Even *Gnamptogenys moelleri* (Forel, 1912), a species with differing chromosome numbers between two populations (2n=34 and 44), has a single intrachromosomal rDNA site. *Gnamptogenys tortuolosa* is an exception, with rDNA clusters occurring over the entire long arm (this study). In the genus *Anochetus*, the chromosome number ranges from 2n=30 to 46, and a single pericentromeric rDNA site has been observed (Santos *et al.*, 2010).

All analyzed fungus-farming ants (Attina) had a single rDNA site located in the intrachromosomal region (Table 2). In the genus *Mycetophylax*, although chromosome number differs among species (2n=26 to 36), a single rDNA site is located in the pericentromeric or terminal region in all the species of the genus (Micolino *et al.*, 2019a). The leaf-cutting ants are considered the most derived among the Attina species (Schultz and Brady, 2008). *Atta* spp. (2n=22) have a single rDNA site in the interstitial region and *Acromyrmex* spp. (2n=38) have a single rDNA site in the terminal region (Barros *et al.*, 2015, 2016; Teixeira *et al.*, 2017). *Acromyrmex striatus* (Roger, 1863), a sister group of the leaf-cutting ants, has 2n=22 and pericentromeric rDNA clusters that are not located on the same chromosome pair relative to that in *Atta* spp. (Cristiano *et al.*, 2013; Teixeira *et al.*, 2017). In *A. echinatior* (2n=38), rDNA clusters are located in the interstitial region of the same pair as observed in other *Acromyrmex* spp. (this study).

The chromosome number of the Australian bulldog ants is highly variable, ranging from 2n=2 to 76 (Imai *et al.*, 1994). In addition, *Myrmecia* spp. present remarkable patterns of multiple 28S rDNA clusters that are highly dispersed throughout their genomes. The number of rDNA sites increases with the chromosome number of the species. This suggests several ribosomal gene amplification events have occurred in the different species of *Myrmecia* and that they have accumulated in karyotypes throughout the evolution of the genus. Only four out of 16 species from this monophyletic genus
1 have the entire arm or intrachromosomal rDNA clusters restricted to a single pair of
2 chromosomes (Hirai *et al.*, 1994, 1996; Hirai, 2020). This pattern is observed in species
3 with small chromosome numbers, which suggests that a single NOR is plesiomorphic
4 among *Myrmecia* (Hirai, 2020).

5 Carpenter ants (*Camponotus*) from the subgenus *Myrmothrix* have 2n=40
6 chromosomes, with its studied species having a single rDNA site in the terminal region.
7 *Camponotus renggeri* is the only exception, having an additional rDNA cluster at the
terminal region of a medium-sized subtelocentric pair (Aguiar *et al.*, 2017).

9 In the giant ants of the genus *Dinoponera*, two contrasting patterns have been
10 observed: *D. gigantea* (2n=82) has multiple rDNA sites located on its short chromosome
11 arms (Aguiar *et al.*, 2011), whereas *Dinoponera lucida* Emery, 1901 has a higher
12 chromosome number (2n=120), but only a single rDNA site restricted to the
13 intrachromosomal region of its largest chromosome pair (Mariano *et al.*, 2008).

14 Homeology patterns among chromosomal pairs bearing ribosomal genes can be
15 detected in a few ant genera, such as *Gnamptogenys* (*striatula* group), *Camponotus*
16 (*Myrmothrix*), *Atta*, and *Acromyrmex* (Table 2). However, it is speculative to infer such
17 homeology patterns for the entire Formicidae family. Ants constitute an ultra-diverse
18 monophyletic group with more than 13,800 described species (Bolton, 2020). They show
19 a wide range of karyotype variation both in number (2n=2 to 2n=120) and chromosomal
20 morphology (reviewed by Lorite and Palomeque, 2010; Mariano *et al.*, 2019).

2 - Insights concerning the organizational patterns of ribosomal gene clusters in the
ant genome

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The mapping of the ribosomal gene clusters of 63 species distributed in 19 genera and six subfamilies, together with information on their phylogenetic relationships, demonstrated that a single pair of chromosomes bearing the GC-rich rDNA clusters is the most frequent trait among the studied species, regardless of the chromosome number (Table 1, Fig. 4A). Genomes carrying rDNA clusters in more than a single chromosome pair have been observed in non-related taxa, such as *D. gigantea* (Aguiar *et al.*, 2011), *C. renggeri* (Aguiar *et al.*, 2017), and *Myrmecia* spp. (Hirai *et al.*, 1994, 1996). We hypothesize that having a single rDNA site should be considered a plesiomorphic trait because multiple rDNA sites were observed in different non-related lineages that do not share exclusive common ancestry and appear *de novo* throughout the Formicidae family.
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**Figure 4.** Summary of available molecular cytogenetic data (this study and in the literature) concerning rDNA genes in ant species: (A) number of species with single and multiple rDNA sites in a karyotype, (B) location of rDNA genes in chromosomes of ant species, considering the following chromosomal regions: c, centromeric region; p, pericentromeric region; i, interstitial region; t, terminal region; wsa, whole short arm; wla,
1 whole long arm, and (C) number of species with single and multiple rDNA sites by
2 subfamilies of Formicidae.

3 In eukaryotes, it is common to observe variations in the number of rDNA clusters
4 and the location of these genes in the chromosomes within genera (Sánchez-Gea et al.,
5 2000; Gross et al., 2010; Cabral-de-Mello et al., 2011; Gokhman et al., 2014; Mazzoleni
6 et al., 2018), among populations (Panzera et al., 2014; Ferreti et al., 2019; Menezes et
7 al., 2019), and sexes of the same species (Nakayama et al., 2001; Šťáhlavský et al., 2018).
8 The possession of terminal rDNA clusters seems to be a common trait among mammals,
9 fish, and mollusks, but less so in arthropods (Sochorová et al., 2018). Within the ultra-
10 diverse insect group, the location of rDNA clusters may follow distinct patterns in its two
11 largest orders; terminal rDNA sites are more abundant in Coleoptera, whereas
12 pericentromeric rDNA clusters are more frequent in Orthoptera (Sochorová et al., 2018).
13 In Formicidae, terminal rDNA sites are a less common feature and all species with
14 multiple rDNA clusters show these genes in the entire short chromosome arms including
15 terminal/subterminal regions, such as C. renggeri, D. gigantea, and Myrmecia spp.
16 (Aguiar et al., 2011, 2017; Hirai et al., 1994, 1996).

17 Different hypotheses have been proposed to explain the cytogenetic pattern
18 (conservative or variable) of these rDNA clusters in the genome of several organisms.
19 Such hypotheses are linked to the specific locations of these rDNA clusters on the
20 chromosomes. Rearrangements, such as translocations, unequal exchange, and ectopic
21 recombination mechanisms (i.e., between non-homologous chromosomes), which can
22 lead to gene dispersion or increases in number in the genome, are more likely in the
23 terminal/subterminal regions of chromosomes and are uncommon in the
24 intrachromosomal regions (Martins and Wasko, 2004; Mantovani et al., 2005; Nguyen et
25 al., 2010; Roy and Guerra, 2012; Hirai, 2020).
Based on rDNA mapping data in fish karyotypes, Martins and Wasko (2004) proposed that translocations are more likely to occur in telomeric regions because of their proximity within the interphase nucleus, which originates from the ordering of chromosomes according to Rabl's model. Effects due to the location of ribosomal genes in relation to their dispersion in the karyotype were also observed in Coleoptera (Dutrillaux et al., 2016) and primates (Gerbault-Seureau et al., 2017). These authors argued that translocations in the interstitial position could result in abnormal meiosis and, therefore, unbalanced gametes. In contrast, translocations in terminal positions may increase the number of rDNA genes in the genome. This would lead to fewer meiotic abnormalities and highlights the selection for interstitial rDNA site stability (Dutrillaux et al., 2016; Gerbault-Seureau et al., 2017).

Ectopic recombination is another mechanism suggested to explain the rDNA patterning in moths and butterflies (Nguyen et al., 2010) and plants (Roa and Guerra, 2012). It is also included in the recent model proposed by Hirai (2020). In this model, two mechanisms are important: the “site effect” and the “molecular effect.” The former allows terminal region associations due to the proximity of these regions in a meiotic bouquet. The “site effect” is a precondition for the “molecular effect,” which refers to systems of affinity/non-affinity due to the similarity between rDNA sequences with other repetitive sequences. Thus, rDNA clusters in the terminal regions tend to associate with other repetitive sequences of non-homologous chromosomes more easily, facilitating the occurrence of ectopic recombination and dispersion of these genes in the genome (Hirai, 2020).

There are reports of species with multiple rDNA clusters associated with the centromeres of acrocentric chromosomes (Cazaux et al., 2011). In the recent model proposed by Hirai (2020), the centromeric region of acrocentric chromosomes
1 (chromosomes with a short and heterochromatic arm) that have rDNA genes associated
2 with the centromere may behave as subterminal regions. Therefore, such an arrangement
3 would also facilitate eventual associations of rDNA genes with other repetitive sequences
4 and the occurrence of ectopic recombination, which leads to their dispersal in the genome
5 (for details, see Hirai, 2020).

6 The ant rDNA chromosome evolution seems to be in accordance with the above-
7 mentioned hypothesis about dispersal and NOR location because the single rDNA
8 clusters of most studied species are interstitial or pericentromeric (Fig. 4B). In ant species,
9 terminal rDNA clusters are prone to rearrangements that lead to their dispersal.
10 *Camponotus renggeri* (Aguiar *et al.*, 2017), *D. gigantea* (Aguiar *et al.*, 2011), and
11 *Myrmecia* spp. present multiple NORs in the entire short chromosome arms including
12 terminal/subterminal regions, which facilitates the association of these genes with the
13 heterochromatic sequences of other non-homologous acrocentric chromosomes during
14 meiosis and the subsequent occurrence of ectopic recombination (Hirai, 2020). This
15 pattern may be applicable to different ant groups. In addition, inversions have been shown
16 to change the position of rDNA genes in *A. echinatior* (this study), *Dolichoderus* spp.
17 (Santos *et al.*, 2016), and *Myrmecia* spp. (Hirai *et al.*, 1996).
<figure><img src="image_9.png" /><figcaption>**Figure 5.** Summary of available data (this study and literature) concerning the number and position of rDNA genes in ant species, along with their degree of relatedness and diploid chromosome numbers. Ideograms of rDNA-bearing chromosomes show number</figcaption></figure>
and location (terminal, interstitial, pericentromeric or centromeric) of rDNA clusters (red) in the haploid complement. Phylogenetic relationships are based on 1- Moreau and Bell (2013), 2 - Schmidt (2013), 3 - Larabee et al. (2016), 4 - Santos et al. (2016), 5 - Hasegawa and Crozier (2006), 6 - Ward et al. (2015), 7 - Solomon et al. (2019), 8 - Micolino et al. (2019a), 9 - Bacci et al. (2009), 10 - Queiroz (2015). Colours of phylogenetic branches indicate the following subfamilies: red, Ponerinae; purple, Dolichoderinae; pink, Myrmeciinae; blue, Formicinae; green, Myrmicinae; gray, Ectatomminae.

A single rDNA site located in the terminal region or entire chromosomal arm was observed in some ant species. The repetitive sequences in the subterminal/terminal chromosome regions probably do not form affinity systems with ribosomal genes (the so-called molecular effect; for details, see Hirai, 2020) in these species. Therefore, rDNA clusters are restricted to a single chromosomal pair. Future studies focusing on the characterization of repetitive sequences that make up the heterochromatin of these species will help clarify this hypothesis.

Size heteromorphisms are frequent in karyotypes where the rDNA clusters have terminal positions in the chromosomes, as reported in this study (G. tortuolosa and C. atriceps) as well as in other ants (Aguiar et al., 2017) and insects in general (Cabral-de-Mello et al., 2011; Maryańska-Nadachowska et al., 2016; Andrade-Souza et al., 2018). Subtle variations in the size of the rDNA clusters between homologous chromosomes can be observed as a result of late condensation during cell division (Sumner, 2003). However, large variations, such as those mentioned above, at the terminal region on the chromosome are usually related to duplications/deletions as a result of unequal exchange (Schubert and Lysack, 2011). It is believed that exchanges are less common in intrachromosomal regions (Hirai, 2020).
<table><caption>1 **Table 1.** Summary of the available molecular cytogenetic data concerning major ribosomal genes (45S, 28S or 18S) detected by FISH in ants</caption><thead><tr><th>Ant species</th><th>2n/(n)</th><th>Location of rDNA genes in karyotype</th><th>Ideogram</th><th>Co-localization CMA<sub>3</sub>/rDNA</th><th>Reference CMA<sub>3</sub>/rDNA</th></tr></thead><tbody><tr><td colspan="6" style="text-align: center; font-weight: bold;">Neotropical ants</td></tr><tr><td colspan="6" style="text-align: left; font-weight: bold;">Subfamily Myrmicinae</td></tr><tr><td>Acromyrmex aspersus</td><td>38</td><td>Largest subtelocentric pair</td><td></td><td>Yes</td><td>Teixeira et al. (2017)</td></tr><tr><td>Acromyrmex coronatus</td><td>38</td><td>Largest subtelocentric pair</td><td></td><td>Yes</td><td>Barros et al. (2016)</td></tr><tr><td>Acromyrmex disciger</td><td>38</td><td>Largest subtelocentric pair</td><td></td><td>Yes</td><td>Barros et al. (2016)</td></tr><tr><td>Acromyrmex echination</td><td>38</td><td>Largest subtelocentric pair</td><td></td><td>Yes</td><td>Barros et al. (2016)/<br>Present study</td></tr><tr><td>Acromyrmex niger</td><td>38</td><td>Largest subtelocentric pair</td><td></td><td>Yes<sup>a</sup></td><td>Barros et al. (2016)</td></tr><tr><td>Acromyrmex striatus</td><td>22</td><td>2nd metacentric pair</td><td></td><td>Yes<sup>a</sup></td><td>Cristiano et al. (2013)/<br>Teixeira et al. (2017)</td></tr><tr><td>Acromyrmex subterraneus<br>molestans</td><td>38</td><td>Largest subtelocentric pair</td><td></td><td>Yes</td><td>Teixeira et al. (2017)</td></tr><tr><td>Atta bisphaerica</td><td>22</td><td>4th metacentric pair</td><td></td><td>Yes</td><td>Barros et al. (2014)/<br>Teixeira et al. (2017)</td></tr><tr><td>Atta laevigata</td><td>22</td><td>4th metacentric pair</td><td></td><td>Yes</td><td>Barros et al. (2014)/<br>Teixeira et al. (2017)</td></tr><tr><td>Atta robusta</td><td>22</td><td>4th metacentric pair</td><td></td><td>Yes</td><td>Barros et al. (2015)</td></tr><tr><td>Atta sexdens rubropilosa</td><td>22</td><td>4th metacentric pair</td><td></td><td>Yes</td><td>Barros et al. (2014)/<br>Teixeira et al. (2017)</td></tr></tbody></table>
<table><caption>Table 1. Cont.</caption><thead><tr><th>Ant species</th><th>2n/(n)</th><th>Localization of rDNA genes in karyotype</th><th>Ideogram</th><th>Co-localization CMA3/rDNA</th><th>Reference CMA3/rDNA</th></tr></thead><tbody><tr><td>Crematogaster longispina</td><td>24</td><td>Largest metacentric pair</td><td>□X□</td><td>Yes</td><td>Present study</td></tr><tr><td>Mycetophylax conformis</td><td>30</td><td>11th metacentric pair</td><td>□X□</td><td>Yes</td><td>Cardoso et al. (2014)/<br/>Micolino et al. (2019a)</td></tr><tr><td>Mycetophylax morschi</td><td>26</td><td>2nd submetacentric pair</td><td>□X□</td><td>–</td><td>Micolino et al. (2019a)</td></tr><tr><td>Mycetophylax morschi</td><td>28</td><td>7th metacentric pair</td><td>□X□</td><td>–</td><td>Micolino et al. (2019a)</td></tr><tr><td>Mycetophylax morschi</td><td>30</td><td>Acrocentric pair</td><td>X□□</td><td>–</td><td>Micolino et al. (2019a)</td></tr><tr><td>Mycetophylax simplex</td><td>36</td><td>Smallest metacentric pair</td><td>□X□</td><td>Yes</td><td>Cardoso et al. (2014)/<br/>Micolino et al. (2019a)</td></tr><tr><td>Mycocepurus goeldii</td><td>8</td><td>2nd metacentric pair</td><td>□X□</td><td>Yes<sup>a</sup></td><td>Barros et al. (2010)/<br/>Barros et al. (2012)</td></tr><tr><td>Myrmicocrypta sp.</td><td>30</td><td>9th metacentric pair</td><td>□X□</td><td>Yes</td><td>Present study</td></tr><tr><td>Pheidole germaini</td><td>22</td><td>Subtelocentric pair</td><td>X□□</td><td>Yes</td><td>Present study</td></tr><tr><td>Solenopsis geminata</td><td>32</td><td>Smallest submetacentric pair</td><td>□X□</td><td>Yes</td><td>Present study</td></tr><tr><td>Strumigenys diabola</td><td>40</td><td>4th submetacentric pair</td><td>X□□</td><td>Yes</td><td>Present study</td></tr><tr><td>Trachymyrmex holmgreni</td><td>20</td><td>4th metacentric pair</td><td>□X□</td><td>Yes</td><td>Barros et al. (2018)</td></tr><tr><td colspan="6"><strong>Subfamily Formicinae</strong></td></tr><tr><td>Camponotus atriceps<sup>c</sup></td><td>40</td><td>2nd submetacentric pair</td><td>□X□</td><td>Yes</td><td>Present study</td></tr></tbody></table>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>

<figure><img src="image_15.png" /></figure>

<figure><img src="image_16.png" /></figure>

<figure><img src="image_17.png" /></figure>

<figure><img src="image_18.png" /></figure>

<figure><img src="image_19.png" /></figure>

<figure><img src="image_20.png" /></figure>

<figure><img src="image_21.png" /></figure>

<figure><img src="image_22.png" /></figure>

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<table><caption>1 Table 1. Cont.</caption><thead><tr><th>Ant species</th><th>2n/(n)</th><th>Localization of rDNA genes in karyotype</th><th>Ideogram</th><th>Co-localization CMA3/rDNA</th><th>Reference CMA3/rDNA</th></tr></thead><tbody><tr><td>Camponotus cingulatus<sup>c</sup></td><td>40</td><td>2nd submetacentric pair</td><td>□</td><td>Yes</td><td>Aguiar et al. (2017)</td></tr><tr><td>Camponotus renggeri</td><td>40</td><td>2nd submetacentric pair and medium-sized subtelocentric pair</td><td>□</td><td>Yes</td><td>Aguiar et al. (2017)</td></tr><tr><td>Camponotus rufipes<sup>c</sup></td><td>40</td><td>2nd submetacentric pair</td><td>□</td><td>Yes</td><td>Aguiar et al. (2017)</td></tr><tr><td>Gigantiops destructor</td><td>(39)</td><td>8th metacentric pair</td><td>□</td><td>Yes<sup>a</sup></td><td>Present study</td></tr><tr><td><strong>Subfamily Ponerinae</strong></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Anochetus altisquamis<sup>c</sup></td><td>30</td><td>3rd submetacentric pair</td><td>□</td><td>—</td><td>Santos et al. (2010)</td></tr><tr><td>Anochetus horridus<sup>c</sup></td><td>46</td><td>4th telocentric pair</td><td>□</td><td>—</td><td>Santos et al. (2010)</td></tr><tr><td>Anochetus targionii</td><td>30</td><td>7th metacentric pair</td><td>□</td><td>Yes</td><td>Present study</td></tr><tr><td>Dinoponera gigantea</td><td>82</td><td>Multiple pairs</td><td>-</td><td>—</td><td>Aguiar et al. (2011)</td></tr><tr><td>Dinoponera lucida</td><td>(59)</td><td>Largest pair (A<sup>Mt</sup>)</td><td>□</td><td>Yes</td><td>Mariano et al. (2008)</td></tr><tr><td>Odontomachus bauri<sup>d</sup></td><td>44/(22)</td><td>2nd subtelocentric pair</td><td>□</td><td>Yes</td><td>Present study</td></tr><tr><td>Odontomachus haematodus</td><td>44</td><td>3rd subtelocentric pair</td><td>□</td><td>Yes</td><td>Present study</td></tr><tr><td>Pseudoponera gilberti</td><td>22</td><td>Largest metacentric pair</td><td>□</td><td>Yes<sup>a</sup></td><td>Present study</td></tr></tbody></table>
<table><caption>1 Table 1. Cont.</caption><thead><tr><th>Ant species</th><th>2n/(n)</th><th>Localization of rDNA genes in karyotype</th><th>Ideogram</th><th>Co-localization CMA<sub>3</sub>/rDNA</th><th>Reference CMA<sub>3</sub>/rDNA</th></tr></thead><tbody><tr><td colspan="6"><strong>Subfamily Ectatomminae</strong></td></tr><tr><td>Gnamptogenys moelleri</td><td>34, 44</td><td>4th metacentric pair</td><td></td><td>Yes</td><td>Teixeira et al. (2020)</td></tr><tr><td>Gnamptongeys regularis<sup>d</sup></td><td>26</td><td>Metacentric and submetacentric homologous</td><td></td><td>Yes</td><td>Teixeira et al. (2020)</td></tr><tr><td>Gnamptogenys striatula</td><td>32</td><td>5th metacentric pair</td><td></td><td>Yes</td><td>Teixeira et al. (2020)</td></tr><tr><td>Gnamptogenys striatula</td><td>34</td><td>4th metacentric pair</td><td></td><td>Yes</td><td>Teixeira et al. (2020)</td></tr><tr><td>Gnamptogenys triangularis</td><td>24</td><td>Largest metacentric pair</td><td></td><td>Yes</td><td>Teixeira et al. (2020)</td></tr><tr><td>Gnamptogenys tortuolosa<sup>c</sup></td><td>44</td><td>Subtelocentric and submetacentric homologous</td><td></td><td>Yes</td><td><strong>Present study</strong></td></tr><tr><td colspan="6"><strong>Subfamily Dolichoderinae</strong></td></tr><tr><td>Dolichoderus attelaboides</td><td>58</td><td>Largest submetacentric pair</td><td></td><td>Yes</td><td>Santos et al. (2016)</td></tr><tr><td>Dolichoderus bidens</td><td>18</td><td>Largest metacentric pair</td><td></td><td>Yes<sup>a</sup></td><td>Santos et al. (2016)</td></tr><tr><td>Dolichoderus decollatus</td><td>38</td><td>2nd metacentric pair</td><td></td><td>Yes</td><td>Santos et al. (2016)</td></tr><tr><td>Dolichoderus diversus</td><td>22</td><td>Largest metacentric pair</td><td></td><td>Yes<sup>a</sup></td><td>Santos et al. (2016)</td></tr></tbody></table>
<table><caption>1 Table 1. Cont.</caption><thead><tr><th>Ant species</th><th>2n/(n)</th><th>Localization of rDNA genes in karyotype</th><th>Ideogram</th><th>Co-localization CMA<sub>3</sub>/rDNA</th><th>Reference CMA<sub>3</sub>/rDNA</th></tr></thead><tbody><tr><td>Dolichoderus imitator</td><td>38</td><td>Largest metacentric pair</td><td></td><td>Yes</td><td>Santos et al. (2016)</td></tr><tr><td>Dolichoderus lutosus</td><td>10</td><td>2nd metacentric pair</td><td></td><td>Yes<sup>a</sup></td><td>Santos et al. (2016)</td></tr><tr><td>Dolichoderus voraginosus</td><td>20</td><td>Largest metacentric pair</td><td></td><td>No<sup>a</sup></td><td>Santos et al. (2016)</td></tr><tr><td colspan="6" style="text-align: center; font-weight: bold;">Australian ants</td></tr><tr><td colspan="6" style="text-align: left; font-weight: bold;">Subfamily Myrmeciinae</td></tr><tr><td>Myrmecia banksi<sup>c</sup></td><td>10</td><td>Smallest acrocentric pair</td><td></td><td>-</td><td>Hirai et al. (1994), (1996)</td></tr><tr><td>Myrmecia chasei</td><td>47</td><td>Multiple pairs</td><td></td><td>-</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia croslandi<sup>d</sup></td><td>2, 3, 4<sup>b</sup></td><td>Acrocentric pair</td><td></td><td>-</td><td>Hirai et al. (1994), (1996)</td></tr><tr><td>Myrmecia forficata</td><td>52<sup>b</sup>, 54</td><td>Multiple pairs</td><td></td><td>-</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia fulvipes</td><td>48, 50</td><td>Multiple pairs</td><td></td><td>-</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia gulosa</td><td>38<sup>b</sup></td><td>Multiple pairs</td><td></td><td>-</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia haskinsorum</td><td>18</td><td>Multiple pairs</td><td></td><td>-</td><td>Hirai et al. (1994), (1996)</td></tr></tbody></table>

2
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<caption>Table 1. Cont.</caption>

2

<table><thead><tr><th>Ant species</th><th>2n/(n)</th><th>Localization of rDNA genes in karyotype</th><th>Ideogram</th><th>Co-localization CMA<sub>3</sub>/rDNA</th><th>Reference CMA<sub>3</sub>/rDNA</th></tr></thead><tbody><tr><td>Myrmecia imaii<sup>c</sup></td><td>8</td><td>Largest acrocentric pair</td><td></td><td>—</td><td>Hirai et al. (1994), (1996)</td></tr><tr><td>Myrmecia mandibularis</td><td>56<sup>b</sup></td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia michaelseni</td><td>27</td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia occidentalis</td><td>64<sup>b</sup></td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia pavida</td><td>44</td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. 1996</td></tr><tr><td>Myrmecia pilosula</td><td>22<sup>b</sup></td><td>Acrocentric pair</td><td></td><td>—</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia pilosula</td><td>19, 23<sup>b</sup>, 27, 32</td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. (1994), (1996)</td></tr><tr><td>Myrmecia simillima</td><td>70<sup>b</sup></td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia arnoldi</td><td>53, 55, 57, 59, 60<sup>b</sup>, 64, 66</td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. (1996)</td></tr><tr><td>Myrmecia vindex</td><td>74, 76</td><td>Multiple pairs</td><td>-</td><td>—</td><td>Hirai et al. (1996)</td></tr></tbody></table>

## Palearctic ant

### Subfamily Dolichoderinae

Tapinoma nigerrimum 18

Largest submetacentric pair

<figure><img src="image_23.png" /></figure>

Yes

Lorite et al. (1997)

3

4

<footnote>a: Additional markers CMA<sub>3</sub><sup>+</sup>. b: Data showed by Hirai et al. (1996). c: Heteromorphism of rDNA clusters between the homologous.</footnote>

<footnote>d: Polymorphism of rDNA clusters between the homologous. – unavailable data.</footnote>
1
2

<table><caption>**Table 2.** The relationship between Neotropical ant species studied using 18S rDNA FISH and their chromosome number</caption><thead><tr><th>Ant species</th><th>Subfamily</th><th>Locality</th><th>2n</th><th>3</th></tr></thead><tbody><tr><td>Acromyrmex echinatior (Forel, 1899)</td><td>Myrmicinae</td><td>Barro Colorado – Panamá</td><td>38</td><td>4</td></tr><tr><td>Anochetus targionii Emery 1894</td><td>Ponerinae</td><td>Campus Agronomique, Kourou – FG</td><td>30</td><td>5</td></tr><tr><td>Camponotus atriceps (Smith, 1858)</td><td>Formicinae</td><td>Viçosa – Minas Gerais – BR</td><td>40</td><td>6</td></tr><tr><td>Crematogaster longispina Emery, 1890</td><td>Myrmicinae</td><td>La Montagne des Singes – FG</td><td>24</td><td>7</td></tr><tr><td>Gigantiops destructor (Fabricius, 1804)</td><td>Formicinae</td><td>Sinnamary – FG</td><td>78</td><td>8</td></tr><tr><td>Gnamptogenys tortuolosa (Smith, 1858)</td><td>Ectatominae</td><td>Sinnamary – FG</td><td>44</td><td>10</td></tr><tr><td>Myrmicocrypta sp.</td><td>Myrmicinae</td><td>Sinnamary – FG</td><td>30</td><td>11</td></tr><tr><td>Odontomachus bauri Emery, 1892</td><td>Ponerinae</td><td>Açailândia – Maranhão – BR</td><td>44</td><td>12</td></tr><tr><td>Odontomachus haematodus (Linnaeus, 1758)</td><td>Ponerinae</td><td>Campus Agronomique, Kourou – FG<br>Ubá – Minas Gerais – BR</td><td>44</td><td>13<br>14</td></tr><tr><td>Pheidole germaini Emery, 1896</td><td>Myrmicinae</td><td>Viçosa – Minas Gerais – BR</td><td>22</td><td>15</td></tr><tr><td>Pseudoponera gilberti (Kempf, 1960)</td><td>Ponerinae</td><td>Sinnamary – FG</td><td>12</td><td>16<br>17</td></tr><tr><td>Solenopsis geminata (Fabricius, 1804)</td><td>Myrmicinae</td><td>Sinnamary – FG</td><td>32</td><td>18</td></tr><tr><td>Strumigenys diabola Bolton 2000</td><td>Myrmicinae</td><td>Sinnamary – FG</td><td>40</td><td>19</td></tr><tr><td></td><td></td><td></td><td></td><td>20</td></tr></tbody></table>

21

FG: French Guiana, BR: Brazil.
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Size variations in the rDNA clusters can be observed when these genes are located
in the interstitial/pericentromeric region of the chromosomes, as seen in *Gnamptogenys
regularis* Mayr, 1870 (Teixeira *et al.*, 2020) and in *O. bauri* (this study). A different path
seems to be involved in the evolution of these karyotypes compared to the rearrangements
involved in terminal rDNA heteromorphisms. In these cases, the mechanism may be
associated with the formation of extrachromosomal circular DNA (eccDNA), which is
likely to form tandem repetitive sequences similar to the rDNA genes (Cohen and Segal,
2009). These eccDNAs may be lost, leading to deletions in the original rDNA sequences,
or they may be replicated via a rolling circle mechanism and reintegrated into the original
chromosome, producing duplications of these repetitive sequences (Cohen and Segal,
2009).

In ants as well as in other eukaryotes, rDNA clusters are located in GC-rich
regions (Symonová, 2019) and, therefore, usually coincide with CMA3+ bands, possibly
as a result of GC-biased gene conversion (gBGC) over the course of evolutionary time
(Escobar et al., 2011). This co-localization of GC-rich regions and rDNA was observed
for all the ants studied to date (Table 2), with the exception of *D. voraginosus* (Santos et
al., 2016). However, GC-rich chromatin is not always an indication of ribosomal genes,
as seen here in *G. destructor*, *P. gilberti*, and in some *Dolichoderus* spp. and fungus-
farming ants (Table 2).

In insects, a compilation of previous data concerning rDNA genes has been
conducted, for example, in moths and butterflies (Lepidoptera; Nguyen *et al.*, 2010),
beetles (Coleoptera; Dutrillaux and Dutrillaux, 2012), and kissing bugs (Heteroptera;
Panzera *et al.*, 2012). However, this is the first survey of Hymenoptera species. We have
compiled available information and new data on 13 Neotropical ant species. In different
organisms, including ants, the number and location of chromosomes bearing rDNA
clusters within the genome follow general patterns that govern the modes of evolution for

these genes (Martins and Wasko, 2004; Nguyen et al., 2010; Dutrillaux et al., 2016;

Gerbault-Seureau et al., 2017; Hirai, 2020; this study). We can conclude that having only

a single pair of chromosomes bearing rDNA clusters is more common in the ant genome

because of the pericentromeric/interstitial location of these genes on the chromosomes.

Intrachromosomal regions are sites with low frequencies of rearrangements, such as non-

Robertsonian translocations and ectopic recombination, and are therefore less prone to

meiotic abnormalities. It should be assumed that the chromosomal location of rDNA

clusters influences the dispersion of these genes within the karyotype.

Future studies will allow the mapping of rDNA genes in more ant taxa, including

the other remaining subfamilies. Other repetitive sequences, such as 5S rDNA and histone

genes, may also be mapped in ant species as a tool to investigate further patterns that

reflect the relationship between chromosomal location and dispersion in the genome.

Finally, a solid understanding of the evolutionary patterns of ribosomal gene dispersal in

ant chromosomes may provide a comparative model for other insects.

16

## 17 Experimental Procedures

18

## 1 - Obtaining samples for analysis

20

Field surveys to collect ant colonies were performed in French Guiana, Brazil, and

Panama (Table 1) from the following locations: *La Montagne des Singes*, Kourou

(5.07225, -52.69407), Campus Agronomique, Kourou (5.17312, -52.65480), and

Sinnamary (5.28482, -52.91403), all in French Guiana; Viçosa (-20.757041, -42.873516)
and Ubá (-21.128880, -42.937646), both in Minas Gerais, Brazil, and Açailândia (-4.84200, -47.29667) in Maranhão, Brazil; and Barro Colorado Island (9.150000, -79.83333) in Panama. Sampling permit in Brazil was provided by the Instituto Chico Mendes de Conservação da Biodiversidade (ICMBio) (SISBIO accession numbers 32459 and 62598). Adult specimens were identified by Dr. Jacques H. C. Delabie and deposited in the ant collection at the Laboratório de Mirmecologia do Centro de Pesquisas do Cacau (CPDC/Brazil), in Bahia, Brazil, under records #5802, #5803, and #5804.

In this study, previous molecular cytogenetic data related to rDNA genes (45S, 18S, or 28S) mapped by FISH from 50 ant species were used. For comparative analysis, the following traits were considered for each species: chromosome number, number of rDNA-bearing chromosomes, location of rDNA clusters in the karyotype, and co-localization of CMA₃ fluorochrome and rDNA clusters. Ag-NOR data were disregarded due to the highly variable patterns and unreliable results (for details, see Introduction).

## 2 – Chromosome preparation

Mitotic metaphase chromosomes were obtained from the cerebral ganglia of larvae after meconium elimination, using colchicine hypotonic solution (0,005%) and fixatives according to the methods described by Imai *et al.* (1988).

## 3 - Staining with fluorochromes chromomycin A₃ (CMA₃) and 4'-6-diamidino-2-phenylindole (DAPI)

Metaphase chromosomes of all the species, except *Ac. echinatior*, were stained with the fluorochromes CMA₃ and DAPI for the detection of GC and AT-rich regions,
respectively, based on the technique proposed by Schweizer (1980). The *Ac. echination*
samples studied here correspond to the same colonies studied by Barros *et al.* (2016) and
the CMA<sub>3</sub>/DAPI staining in this species was performed by these authors.

4

5 **4 - Fluorescent *in situ* hybridization with rDNA 18S probe**

6

The 18S rDNA probes were obtained by amplification via polymerase chain
reaction (PCR) using *Melipona quiquefasciata* Lepeletier, 1836, rDNA primers 18SF1
(5'-GTC ATA GCT TTG TCT CAA AGA-3') and 18SR1.1 (5'-CGC AAA TGA AAC
TTT AAT CT-3') (Pereira, 2006) in the genomic DNA from the ant *Camponotus rufipes*
(Fabricius, 1775). Gene amplification was performed following Pereira (2006). The
probes were labeled by an indirect method using digoxigenin-11-dUTP (Roche Applied
Science, Mannheim, Germany), and the FISH signals were detected with anti-
digoxigenin-rhodamine (Roche Applied Science), following the manufacturer's protocol.

The rDNA 18S genes were mapped by FISH, following the protocol of Pinkel et
al. (1986). The slides were treated with RNase A (100 µg/mL) and kept in a moist
chamber at 37°C for 1h. After that, they were washed in 2×SSC for 5 min, incubated in
5 µg/ml pepsin in 0.01 N HCl for 10 min, washed in 1×PBS for 5 min and dehydrated in
50%, 70% and 100% alcohol series for 2 min each. After this pretreatment, metaphase
chromosomes were denatured in 70% formamide/2×SSC at 75°C for 3 min, and 20 µL of
hybridization mix including 200 ng of labeled probe, 2×SSC, 50% formamide, and 10%
dextran sulfate was denatured for 10 min at 85°C and added on preparations. The slides
were kept in a moist chamber at 37°C overnight. Then, the slides were washed in 2×SSC
for 5 min; the detection solution including anti-digoxigenin-rhodamine was added on
slides that were kept in a moist chamber at 37°C for 1h. The slides were washed in
1

4×SSC/Tween, and dehydrated in an alcohol series. Finally, counterstaining with DAPI

2

(DAPI Fluoroshield, Sigma Aldrich) was performed.

3

4

## 5 - Chromosomal analysis

5

6

Chromosomes were arranged in order of decreasing size and based on the ratio of the chromosomes arm lengths (r = long arm/short arm), according to the classification proposed by Levan *et al.* (1964). The chromosomes were classified as m = metacentric (r = 1–1.7), sm = submetacentric (r = 1.7–3), st = subtelocentric (r = 3–7), and a = acrocentric (r > 7); they were organized using Adobe Photoshop® 21.1.1 and measured using Image Pro Plus®. Ideograms of the NOR-bearing chromosome/chromosomes (i.e., graphical representation of the chromosomes concerning the rDNA clusters) of the ant species were then designed with the Easy Idio software (Diniz and Xavier, 2006).

7

8

9

10

11

12

13

14

For the fluorochrome staining and FISH 18S rDNA technique, 30 metaphases from at least three individuals of each species were analyzed. In the case of *O. bauri*, which presented a chromosomal polymorphism involving rDNA clusters, seven individuals were analyzed (six females and one male). The metaphases were analyzed and photographed using a fluorescence microscope, Olympus BX60, attached to an image system, QColor Olympus®, with the filters WB (450–480 nm), WU (330-385 nm), and WG (510-550 nm) for the fluorochromes CMA₃, DAPI, and rhodamine, respectively.

15

16

17

18

19

20

21

22

## 6 - Phylogenetic relationships

23

24

The phylogenetic relationship among ant species was determined by associating with previously published molecular phylogenies. The resultant cladogram topology at

25
1 the subfamily level was determined following Moreau and Bell (2013). The Poneroid
2 clade topology was determined according to Schmidt (2013) and Larabee et al. (2016);
3 the clade topology for the subfamily Dolichoderinae was determined according to Santos
4 et al. (2016) and that for the subfamily Myrmicinae was determined according to Bacci
5 et al. (2009), Ward et al. (2015), Queiroz (2015), Solomon et al. (2019), and Micolino et
6 al. (2019a). The topology of the species groups within the subfamily Myrmeciinae was
7 determined according to Hasegawa and Crozier (2006).

8

9 Acknowledgments

10

11 We would like to thank Dr. Cléa S. F. Mariano for kindly providing *Acromyrmex*
12 *echinatior* samples, Dr. Jacques H. C. Delabie for species identification, and Dr. Luiz
13 Fernando Gomes for his valuable assistance in the laboratory with *Odontomachus bauri*
14 chromosome preparations. We also acknowledge Laboratório de Biologia Molecular de
15 Insetos of the Universidade Federal de Viçosa (UFV) for technical support. GAT thanks
16 the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) and
17 Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) for the
18 scholarships granted. Financial support for this study was provided by "Investissement
d'Avenir" grants managed by the French Agence Nationale de la Recherche (DRIHM
19 ref. ANR-11-LABX-0010 and CEBA, ref. ANR-10-LABX-25-01), by the PO-FEDER
20 2014-2020, Région Guyane (BiNG, ref. GY0007194), and by the Programa de Auxílio
21 ao Pesquisador – PAPESQ/UNIFAP/2019.
22
<figure><img src="image_24.png" /><figcaption>**Figure S1.** Staining with chromomycin A₃ showing GC-rich regions (arrows) of Ponerinae ants: (A) *Pseudoponera gilberti* (2n=12), (B) *Anochetus targionii* (2n=30), (C) *Odontomachus haematodus* (2n=44, French Guiana), (D, E) *Odontomachus bauri* showing the homozygous and heterozygous states, respectively (2n=44), and (F) male of *Odontomachus bauri* with the smaller size regions rich in GC base pairs. Bars = 5 µm.</figcaption></figure>
<figure><img src="image_25.png" /><figcaption>**Figure S2.** Staining with chromomycin A₃ showing GC-rich regions (arrows) of Myrmicinae ants: (A) *Pheidole germaini* (2n=22), (B) *Crematogaster longispina* (2n=24), (C) *Solenopsis geminata* (n=16), (D) *Myrmicocrypta* sp. (2n=30), (E) *Strumigenys diabola* (2n=40). Bars = 5 µm.</figcaption></figure>
<figure><img src="image_26.png" /><figcaption>**Figure S3** Staining with chromomycin A₃ showing GC-rich regions (arrows) of an Ectatomminae ant (A) *Gnamptogenys tortuolosa* (2n=44), and Formicinae ants (B) *Camponotus atriceps* (2n=40) and (C, D) *Gigantiops destructor* (2n=78, n=39). Bars = 5 µm.</figcaption></figure>
1 **References**

2

3 Aguiar, H.J.A.C., Barros, L.A.C., Alves, D.R., Mariano, C.S.F., Delabie, J.H.C. and

4 Pompolo, S.G. (2017) Cytogenetic studies on populations of *Camponotus rufipes*

5 (Fabricius, 1775) and *Camponotus renggeri* Emery, 1894 (Formicidae: Formicinae).

6 *PLoS ONE*, **12**(5), e0177702.

7 Aguiar, H.J.A.C., Barros, L.A.C., Mariano, C.S.F., Delabie, J.H.C. and Pompolo, S.G.

8 (2011) 45S rDNA localization for the giant Ant *Dinoponera gigantea* with Evolutionary

9 inferences for the *Dinoponera* Genus (Formicidae: Ponerinae). *Sociobiology*, **57**(3),
10 607–620.

11 Andrade-Souza, V., Duarte, O.M.P., Martins, C.C.C., Santos, I.S., Costa, M.G.C. and

12 Costa, M.A. (2018) Comparative molecular cytogenetics of *Melipona* Illiger species

13 (Hymenoptera: Apidae). *Sociobiology*, **65**(4), 696–705.

14 Bacci, Jr. M., Solomon, S.E., Mueller, U.G., Martins, V.G., Carvalho, A.O.R., Vieira,

15 L.G.E. and Silva-Pinhati, A.C.O. (2009). Phylogeny of leafcutter ants in the genus *Atta*

16 Fabricius (Formicidae: Attini) based on mitochondrial and nuclear DNA sequences.

17 *Molecular Phylogenetics and Evolution*, **51**(3), 427–437.

18 Barbosa, P., Leal, E.V., Silva, M., Almeida, M.C., Moreira-Filho, O. and Artoni, R.F.

19 (2017). Variability and evolutionary implications of repetitive DNA dynamics in

20 genome of *Astyanax scabripinnis* (Teleostei, Characidae). *Comparative Cytogenetics*,
21 **11**(1), 143–162.

22 Barros, L.A.C., Aguiar, H.J.A.C., Andrade-Souza, V., Mariano, C.S.F., Delabie, J.H.C.

23 and Pompolo, S.G. (2012). Occurrence of pre-nucleolar bodies and 45S rDNA location
1 on the chromosomes of the ant *Mycocepurus goeldii* (Forel) (Formicidae, Myrmicinae,
2 Attini). *Hereditas*, **149**, 50–54.

3 Barros, L.A.C., Aguiar, H.J.A.C., Mariano, C.S., Andrade-Souza, V., Costa, M.A.,

4 Delabie, J.H.C. and Pompolo, S.G. (2016). Cytogenetic data on six leafcutter ants of the
genus *Acromyrmex* Mayr, 1865 (Hymenoptera, Formicidae, Myrmicinae): insights into
chromosome evolution and taxonomic implications. *Comparative Cytogenetics*, **10**(2),
229–243.

8 Barros, L.A.C., Aguiar, H.J.A.C., Mariano, C.S.F., Delabie, J.H.C. and Pompolo, S.G.
(2010). Cytogenetic characterization of the lower-Attine *Mycocepurus goeldii*
(Formicidae: Myrmicinae: Attini). *Sociobiology*, **56**, 57–66.

11 Barros, L.A.C., Aguiar, H.J.A.C., Teixeira, G.A., Mariano, C.S.F., Teixeira, M.C.,
12 Delabie, J.H.C. and Pompolo, S.G. (2015). Cytogenetic data on the threatened leafcutter
13 ant *Atta robusta* Borgmeier, 1939 (Formicidae: Myrmicinae: Attini). *Comptes Rendus
14 Biologies*, **338**(10), 660–665.

15 Barros, L.A.C., Mariano, C.S.F., Pompolo, S.G., and Delabie, J.H.C. (2009). Hsc-FA
16 and NOR bandings on chromosomes of the giant ant *Dinoponera lucida* Emery, 1901
17 (Hymenoptera: Formicidae). *Comparative Cytogenetics*, **3**, 97-102.

18 Barros, L.A.C., Teixeira, G.A., Aguiar, H.J.A.C., Lopes, D.M. and Pompolo, S.G.
19 (2018). Cytogenetic studies in *Trachymyrmex holmgreni* Wheeler, 1925 (Formicidae:
20 Myrmicinae) by conventional and molecular methods. *Sociobiology*, **65**(2), 185–190.

21 Barros, L.A.C., Teixeira, G.A., Aguiar, H.J.A.C., Mariano, C.S.F, Delabie, J.H.C. and
22 Pompolo, S.G. (2014). Banding patterns of three leafcutter ant species of the genus *Atta*
23 (Formicidae: Myrmicinae) and Chromosomal Inferences. *Florida Entomologist*, **97**(4),
24 1694–1701.
1 Bolton, B. 2020. An online catalog of the ants of the world. Available from

2 https://antcat.org. (accessed 15th May, 2020)

3 Britton-Davidian, J., Cazaux, B. and Catalan, J. (2012). Chromosomal dynamics of

4 nucleolar organizer regions (NORs) in the house mouse: micro-evolutionary insights.

5 *Heredity*, **108**(1), 68-74.

6 Cabral-de-Mello, D.C., Oliveira, S.G., Moura, R.C. and Martins, C. (2011).

7 Chromosomal organization of the 18S and 5S rRNAs and histone H3 genes in

8 Scarabaeinae coleopterans: insights into the evolutionary dynamics of multigene

9 families and heterochromatin. *BMC Genetics*, **12**, 88.

10 Cardoso, D.C., Pompolo, S.G., Cristiano, M.P. and Tavares, M.G. (2014). The role of

11 fusion in ant chromosome evolution: insights from cytogenetic analysis using a

12 molecular phylogenetic approach in the genus *Mycetophylax*. *PLoS ONE*, **9**(4), e95408.

13 Castro, C.P.M., Cardoso, D.C., Micolino, R., and Cristiano, M.P. (2020). Comparative

14 FISH-mapping of TTAGG telomeric sequences to the chromosomes of leafcutter ants

15 (Formicidae, Myrmicinae): is the insect canonical sequence conserved? *Comparative*

16 *Cytogenetics*, **14**(3), 369-385.

17 Cazaux, B., Catalan, J., Veyrunes, F., Douzery, E.J. and Britton-Davidian, J. (2011).

18 Are ribosomal DNA clusters rearrangement hotspots? A case study in the genus *Mus*

19 (Rodentia, Muridae). *BMC Evolutionary Biology*, **11**(1), 124.

20 Cholak, L.R., Haddad, C.F. and Parise-Maltempi, P.P. (2020). Cytogenetic analysis of

21 the genus *Thoropa* Cope, 1865 (Anura-Cycloramphidae) with evolutionary inferences

22 based on repetitive sequences. *Genetics and Molecular Biology*, **43**(3).
1 Cohen, S. and Segal, D. (2009). Extrachromosomal Circular DNA in Eukaryotes:

2 possible involvement in the plasticity of tandem repeats. *Cytogenetic and Genome Research*, **124**, 327–338.

3

4 Cristiano, M.P., Cardoso, D.C. and Fernandes-Salomão, T.M. (2013). Cytogenetic and molecular analyses reveal a divergence between *Acromyrmex striatus* (Roger, 1863) and other congeneric species: taxonomic implications. *PLoS ONE*, **8**(3), e59784.

7 Cross, I., Vega, L., and Rebordinos, L. (2003). Nucleolar organizing regions in *Crassostrea angulata*: chromosomal location and polymorphism. *Genetica*, **119**(1), 65-74.

10 Degrandi, T.M., Gunski, R.J., Garnero, A.D.V., Oliveira, E.H.C.D., Kretschmer, R.,

11 Souza, M.S.D., Barcellos S.A. and Hass, I. (2020). The distribution of 45S rDNA sites in bird chromosomes suggests multiple evolutionary histories. *Genetics and Molecular Biology*, **43**(2).

14 Diniz, D. and Xavier, P.M. (2006). Easy Idio.

15 Dobigny, G., Ozouf-Costaz, C., Bonillo, C. and Volobouev, V. (2002). “Ag-NORs” are not always true NORs: new evidence in mammals. *Cytogenetic and Genome Research*, **98**(1), 75-77.

18 Dutra, R.T., Bitencourt, J., Netto, M.R.C.B., Paim, F.G., Sarmento-Soares. L.M. and Afonso, P.R.A.M. (2020). Chromosomal markers are useful to species identification in Rivulidae (Cyprinodontiformes, Aplocheiloidei), including the resolution of taxonomic uncertainties in a vulnerable species complex. *Zebrafish*, **17**(1), 48–55.

22 Dutrillaux, A.M. and Dutrillaux, B. (2012). Chromosome analysis of 82 species of Scarabaeoidea (Coleoptera), with special focus on NOR localization. *Cytogenetic and Genome Research*, **136**(3), 208–219.
1 Dutrillaux, A.M., Carton, B., Cacheux, L and Dutrillaux, B. (2016). Interstitial NORs, fragile sites, and chromosome evolution: a not so simple relationship-the example of

2 *Melolontha melolontha* and genus *Protaetia* (Coleoptera: Scarabaeidae). *Cytogenetic and Genome Research*, **149**(4), 304–311.

5 Escobar, J.S., Glémin, S. and Galtier, N. (2011). GC-biased gene conversion impacts ribosomal DNA evolution in vertebrates, angiosperms, and other eukaryotes. *Molecular Biology and Evolution*, **28**(9), 2561–2575.

8 Falcione, C., Hernando, A. and Bressa, M.J. (2018). Comparative cytogenetic analysis in *Erythrolamprus* snakes (Serpentes: Dipsadidae) from Argentina. *Anais da Academia Brasileira de Ciencias*, **90**(2), 1417-1429.

11 Ferreti, A.B.S.M., Ruiz-Ruano, F.J., Milani, D., Loreto, V., Martí, D.A., Ramos, E., Martins, C. and Cabral-de-Mello, D.C. (2019). How dynamic could be the 45S rDNA cistron? An intriguing variability in a grasshopper species revealed by integration of chromosomal and genomic data. *Chromosoma*, **128**, 165–175.

15 Gerbault-Seureau, M., Cacheux, L. and Dutrillaux, B. (2017). The relationship between the (In-) stability of NORs and their chromosomal location: the example of

16 Cercopithecidae and a short review of other primates. *Cytogenetic and Genome Research*, **153**(3), 138–146.

19 Gokhman, V.E., Anokhin, B.A. and Kuznetsova, V.G. (2014). Distribution of 18S rDNA sites and absence of the canonical TTAGG insect telomeric repeat in parasitoid

20 Hymenoptera. *Genetica*, **142**, 317–322.

22 Gokhman, V.E., Bolsheva, N.L., Govind, S. and Muravenko, O.V. (2016). A comparative cytogenetic study of *Drosophila* parasitoids (Hymenoptera, Figitidae)
1 using DNA-binding fluorochromes and FISH with 45S rDNA probe. *Genetica*, **144**(3),

2 335-339.

3 Golub, N.V., Golub, V.B. and Kuznetsova, V.G. (2015). Variability of 18rDNA loci in

4 four lace bug species (Hemiptera, Tingidae) with the same chromosome number.

5 *Comparative Cytogenetics*, **9**(4), 513-522.

6 Gross, M.C., Schneider, C.H., Valente, G.T., Martins, C. and Feldberg E. (2010).

7 Variability of 18S rDNA locus among *Symphysodon* fishes: chromosomal

8 rearrangements. *Journal of Fish Biologies*, **76**(5), 1117-1127.

9 Hasegawa, E, and Crozier, R.H. (2006). Phylogenetic relationships among species

10 groups of the ant genus *Myrmecia*. *Molecular Phylogenetics and Evolution*, **38**, 575-

11 582.

12 Hirai, H. (2020). Chromosome dynamics regulating genomic dispersion and alteration

13 of Nucleolus Organizer Regions (NORs). *Cells*, **9**(4), 971-984.

14 Hirai, H., Yamamoto, M.T., Ogura, K., Satta, Y., Yamada, M., Taylor, R.W. and Imai,

15 H.T. (1994). Multiplication of 28S rDNA and NOR activity in chromosome evolution

16 among ants of the *Myrmecia pilosula* species complex. *Chromosoma*, **103**(3), 171-178.

17 Hirai, H., Yamamoto, M.T., Taylor, R.W. and Imai, H.T. (1996). Genomic dispersion of

18 28S rDNA during karyotypic evolution in the ant genus *Myrmecia* (Formicidae).

19 *Chromosoma*, **105**, 190-196.

20 Howell, W.T., and Black, D.A. (1980). Controlled silver-staining of nucleolus organizer

21 regions with a protective colloidal developer: a 1-step method. *Experientia*, **36**(8),

22 1014-1015.
1 Huang, Y.C., Lee, C.C., Kao, C.Y., Chang, N.C., Lin, C.C., Shoemaker, D. and Wang,

2 J. (2016). Evolution of long centromeres in fire ants. *BMC Evolutionary Biology*, **16**,

3 189.

4 Imai, H., Taylor, R.W., Crosland, M.W. and Crozier, R.H. (1988). Modes of
5 spontaneous chromosomal mutation and karyotype evolution in ants with reference to
6 the minimum interaction hypothesis. *Japanese Journal of Genetics*, **63**, 159–185.

7 Imai, H.T., Hirai, H., Satta, Y., Shiroishi, T., Yamada, M., and Taylor, R.W. (1992).

8 Phase specific Ag-staining of nucleolar organizer regions (NORs) and kinetochores in
9 the Australian ant *Myrmecia croslandi*. *The Japanese Journal of Genetics*, **67**(6), 437-
10 447.

11 Imai, H.T., Taylor, R.W., and Crozier, R.H. (1994). Experimental bases for the
12 minimum interaction theory. I. Chromosome evolution in ants of the *Myrmecia pilosula*
13 species complex (Hymenoptera: Formicidae: Myrmeciinae). *The Japanese Journal of
14 Genetics*, **69**(2), 137-182.

15 Larabee, F.J., Fisher, B.K., Schmidt, C.A., Matos-Maraví, P., Janda, M. and Suarez,
16 A.V. (2016). Molecular phylogenetics and diversification of trap-jaw ants in the genera
17 *Anochetus* and *Odontomachus* (Hymenoptera: Formicidae). *Molecular Phylogenetics
18 and Evolution*, **103**, 143–154.

19 Levsky, J.F. and Singer, R.H. (2003). Fluorescence *in situ* hybridization: past, present
20 and future. *Journal of Cell Science*, **116**, 2833-2838.

21 Liehr, T. (2017). “Classical cytogenetics” is not equal to “banding cytogenetics”.
22 *Molecular Cytogenetics*, **10**, 1–3.

23 Long, E.O. and Dawid, I.B. (1980). Repeated genes in eukaryotes. *Annual Review of
24 Biochemistry*, **49**(1), 727–764.
1 Lorite, P., and Palomeque, T. (2010). Karyotype evolution in ants (Hymenoptera:

2 Formicidae), with a review of the known ant chromosome numbers. *Myrmecological*

3 *News*, **13**(1), 89-102.

4 Lorite, P., Aránega, A.E., Luque, F. and Palomeque, T. (1997). Analysis of the
5 nucleolar organizing regions in the ant *Tapinoma nigerrimum* (Hymenoptera,
6 Formicidae). *Heredity*, **78**, 578-582.

7 Lorite, P., Carrillo, J.A., Tinaut, A. and Palomeque, T. (2004). Evolutionary dynamics
8 of satellite DNA in species of the genus *Formica* (Hymenoptera, Formicidae). *Gene*,
9 **332**, 159-168.

10 Malimpensa, G.D.C., Traldi, J.B., Martinez, J.D.F., Deon, G., Azambuja, M., Nogaroto,
11 V., Vicari M.R. and Moreira-Filho, O. (2020). Chromosomal Diversification in Two
12 Species of *Pimelodus* (Siluriformes: Pimelodidae): Comparative Cytogenetic Mapping
13 of Multigene Families. *Zebrafish*, **17**(4), 278-286.

14 Mantovani, M., Abel, L.D. and Moreira-Filho, O. (2005). Conserved 5S and variable
15 45S rDNA chromosomal localization revealed by FISH in *Astyanax scabripinnis*
16 (Pisces, Characidae). *Genetica*, **123**, 211-216.

17 Mariano, C.S.F., Barros, L.A.C., Velasco, Y.M., Guimarães, I.N., Pompolo, S.G. and
18 Delabie, J.H.C. (2019). Citogenética de hormigas de la región neotropical, In Hormigas
de Colombia (Fernández, F., Guerrero, R. and Delsinne, T., eds.). pp. 131-157.
20 Universidad Nacional de Colombia, Bogotá.

21 Mariano, C.S.F., Pompolo, S.G., Barros, L.A.C., Mariano-Neto, E., Campiolo, S. and
22 Delabie, J.C.H. (2008). A biogeographical study of the threatened ant *Dinoponera
23 lucida* Emery (Hymenoptera: Formicidae: Ponerinae) using a cytogenetic approach.
24 *Insect Conservation and Diversity*, **1**(3), 161-168.
1 Martins, C. and Wasko, A.P. (2004). Organization and evolution of 5S ribosomal DNA

2 in the fish genome. In Focus on genome research (Williams, C.R., ed.). pp. 335–363.

3 Nova Science Publishers Inc, New York.

4 Maryańska-Nadachowska, A., Anokhin, B.A., Gnezdilov, V.M. and Kuznetsova, V.G.

5 2016. Karyotype stability in the family Issidae (Hemiptera, Auchenorrhyncha) revealed

6 by chromosome techniques and FISH with telomeric (TTAGG)n and 18S rDNA probes.

7 *Comparative Cytogenetics*, **10**(3), 347–369.

8 Mazzoleni, S., Rovatsos, M., Schillaci, O. and Dumas, F. (2018). Evolutionary insight

9 on localization of 18S, 28S rDNA genes on homologous chromosomes in Primates

10 genomes. *Comparative Cytogenetics*, **12**(1), 27–40.

11 Menezes, R.S.T., Gazoni, T. and Costa, M.A. (2019). Cytogenetics of warrior wasps

12 (Vespidae: *Synoeca*) reveals intense evolutionary dynamics of ribosomal DNA clusters

13 and an unprecedented number of microchromosomes in Hymenoptera. *Biological*

14 *Jounal of the Linnean Society*, **126**(4), 925–935.

15 Meyne, J., Hirai, H. and Imai, H.T. (1995). FISH analysis of the telomere sequences of

16 bulldog ants (*Mymecia*: Formicidae). *Chromosoma*, **104**, 14–18.

17 Micolino, R., Cristiano, M.P., Travenzoli, N.M., Lopes, D.M. and Cardoso, D.C.

18 (2019a). Chromosomal dynamics in space and time: evolutionary history of

19 *Mycetophylax* ants across past climatic changes in the Brazilian Atlantic coast.

20 *Scientific Reports*, **9**(1), 1–13.

21 Micolino, R., Cristiano, M. P. and Cardoso, D. C. (2019b). Population-based

22 cytogenetic banding analysis and phylogenetic relationships of the neotropical fungus-

23 farming ant *Trachymyrmex holmgreni* Wheeler, 1925. *Cytogenetic and Genome*

24 *Research*, **159**(3), 151-162.
1 Micolino, R., Cristiano, M.P., and Cardoso, D.C. (2020). Karyotype and putative

2 chromosomal inversion suggested by integration of cytogenetic and molecular data of

3 the fungus-farming ant *Mycetomoellerius iheringi* Emery, 1888. *Comparative*

4 *Cytogenetics*, **14**(2), 197-210.

5 Moreau, C.S. and Bell, C.D. (2013). Testing the museum versus cradle tropical

6 biological diversity hypothesis: phylogeny, diversification, and ancestral biogeographic

7 range evolution of the ants. *Evolution*, **67-8**, 2240-2257.

8 Nguyen, P., Sahara, K., Yoshido, A. and Marec, F. (2010). Evolutionary dynamics of

9 rDNA clusters on chromosomes of moths and butterflies (Lepidoptera). *Genetica*, **138**,

10 343-354.

11 Panzera, F., Ferreiro, M.J., Pita, S., Calleros, L., Pérez, R., Basmadjián, Y., Guevara,

12 Y., Brenière, S.F. and Panzera, Y. (2014). Evolutionary and dispersal history of

13 *Triatoma infestans*, main vector of Chagas disease, by chromosomal markers. *Infection*,

14 *Genetics and Evolution*, **27**, 105-113.

15 Panzera, Y.1., Pita, S., Ferreiro, M.J., Ferrandis, I., Lages, C., Pérez, R., Silva, A.E.,

16 Guerra, M. and Panzera, F. (2012). High dynamics of rDNA cluster location in kissing

17 bug holocentric chromosomes (Triatominae, Heteroptera). *Cytogenetics and Genome*

18 *Research*, **38**(1), 56-67.

19 Pereira, J.O.P. (2006). Diversidade genética da abelha sem ferrão *Melipona*

20 *quinquefasciata* baseada no sequenciamento das regiões ITS1 parcial e 18S do DNA

21 ribossômico nuclear. Unpublished D. Phil. Thesis, Universidade Federal do Ceará,

22 Fortaleza, Brazil.

23 Pereira, T.T.P., Reis, A.C.C., Cardoso, D.C. and Cristiano, M.P. (2018). Molecular

24 phylogenetic reconstruction and localization of the (TTAGG)n telomeric repeats in the
1 chromosomes of *Acromyrmex striatus* (Roger, 1863) suggests a lower ancestral

2 karyotype for leafcutter ants (Hymenoptera). *Comparative Cytogenetics*, **12**(1), 13–26.

3 Pinkel, D., Straume, T. and Gray, J.W. (1986). Cytogenetic analysis using quantitative,

4 high-sensitivity, fluorescence hybridization. *Proceedings of the National Academy of Sciences of the United States of America*, **83**, 2934–2938.

6 Queiroz, E.C. (2015). Estudos moleculares em *Acromyrmex* Mayr, 1865. Unpublished

7 D. Phil. Thesis, Universidade Federal do Paraná, Curitiba, Brazil.

8 Roa, F. and Guerra, M. (2012). Distribution of 45S rDNA sites in chromosomes of

9 plants: structural and evolutionary implications. *BMC Evolutionary Biology*, **12**(1), 1-13.

11 Roy, V, Monti-Dedieu, L., Chaminade, N., Siljak-Yakovlev, S., Aulard, S., Lemeunier,

12 F., Montchamp-Moreau, C. (2005). Evolution of the chromosomal location of rDNA
genes in two *Drosophila* species subgroups: *ananassae* and *melanogaster*. *Heredity*,

14 **94**(4), 388–395.

15 Sánchez-Gea, J.F., Serrano, J. and Galián, J. (2000). Variability in rDNA loci in Iberian
species of the genus *Zabrus* (Coleoptera: Carabidae) detected by fluorescence *in situ*
hybridization. *Genome* **43**, 22–28.

18 Santos, I.S., Mariano, C.S.F., Andrade, V., Costa, M.A., Delabie, J.H.C. and Silva, J.G.
(2010). A Cytogenetic Approach to the Study of Neotropical *Odontomachus* and

20 *Anochetus* Ants (Hymenoptera: Formicidae). *Annals of the Entomological Society of America*, **103**(3), 424–429.

22 Santos, I.S., Mariano, C.S.F., Delabie, J.H.C., Costa, M.A., Carvalho, A.F. and Silva, J.G. (2016). "Much more than a neck": karyotype differentiation between *Dolichoderus attelaboides* (Fabricius, 1775) and *Dolichoderus decollatus* F. Smith, 1858
1 (Hymenoptera: Formicidae) and karyotypic diversity of five other Neotropical species
2 of Dolichoderus Lund, 1831. *Myrmecological News*, **23**, 61–69.

3 Schmid, M., Steinlein, C., Feichtinger, W. and Nanda, I. (2017). Chromosome banding in Amphibia. XXXV. Highly mobile nucleolus organizing regions in *Craugastor fitzingeri* (Anura, Craugastoridae). *Cytogenetic and Genome Research*, **152**(4), 180-193.

6 Schmidt, C. (2013). Molecular phylogenetics of ponerine ants (Hymenoptera: Formicidae: Ponerinae). *Zootaxa*, **3647**(2), 201–250.

8 Schubert, I. and Lysack, M.A. (2011). Interpretation of karyotype evolution should consider chromosome structural constraints. *Trends in Genetics*, **27**(6), 207–216.

10 Schultz, T.R. and Brady S.G. (2008). Major evolutionary transitions in ant agriculture.
*Proceedings of the National Academy of Sciences of the United States of America*,
**105**(14), 5435–5440.

13 Schweizer, D. (1980). Simultaneous fluorescent staining of R bands and specific heterocromatic regions (DA/DAPI-bands) in human chromosomes. *Cytogenetics and Cell Genetics*, **27**, 190–193.

16 Sochorová, J., Garcia, S., Gálvez, F., Symonová, R. and Kovařík, A. (2018).
Evolutionary trends in animal ribosomal DNA loci: introduction to a new online database. *Chromosoma*, **127**, 141–150.

19 Solomon, S.E., Rabeling, C., Sosa-Calvo, J., Lopes, C.T., Rodrigues, A., Vasconcelos, H.L., Bacci, Jr.M., Mueller, U.G. and Schultz, T.R. (2019). The molecular phylogenetics of *Trachymyrmex* Forel ants and their fungal cultivars provide insights into the origin and coevolutionary history of ‘higher attine’ ant agriculture. *Systematic Entomology*, **44**(4), 939–956.
1 Šťáhlavský, F., Opatova, V., Just, P., Lotz, L.N. and Haddad, C.R. (2018). Molecular

2 technique reveals high variability of 18S rDNA distribution in harvestmen (Opiliones,

3 Phalangiidae) from South Africa. *Comparative Cytogenetics*, **12**(1), 41–59.

4 Sumner, A.T. (2003). Chromosomes: Organization and Function. Blackwell Publishing,

5 North Berwick, United Kingdom.

6 Symonová, R. (2019). Integrative rDNAomics - Importance of the oldest repetitive

7 fraction of the eukaryote genome. *Genes*, **10**(5), 345–360.

8 Teixeira, G.A., Barros, L.A.C., Aguiar, H.J.A.C. and Pompolo, S.G. (2017).

9 Comparative physical mapping of 18S rDNA in the karyotypes of six leafcutter ant

10 species of the genera *Atta* and *Acromyrmex* (Formicidae: Myrmicinae). *Genetica*, **145*,

11 351–357.

12 Teixeira, G.A., Barros, L.A.C., Lopes, D.M. and Aguiar, H.J.A.C. (2020). Cytogenetic

13 variability in four species of *Gnamptogenys* Roger, 1863 (Formicidae: Ectatomminae)

14 showing chromosomal polymorphisms, species complex, and cryptic species.

15 *Protoplasma*, **257**, 549–560.

16 Vicari, M.R., Artoni, R.F., Moreira-Filho, O. and Bertollo, L.A.C. (2008).

17 Diversification of a ZZ/ZW sex chromosome system in *Characidium* fish (Crenuchidae,

18 Characiformes). *Genetica*, **134**(3), 311.

19 Walker, L.I., Soto, M.A. and Spotorno, Á.E. (2014). Similarities and differences among

20 the chromosomes of the wild guinea pig *Cavia tschudii* and the domestic guinea pig

21 *Cavia porcellus* (Rodentia, Caviidae). *Comparative Cytogenetics*, **8**(2), 153.

22 Ward, P.S., Brady, S.G., Fisher, B.L. and Schultz, T.R. (2015). The evolution of

23 myrmicine ants: phylogeny and biogeography of a hyperdiverse ant clade

24 (Hymenoptera: Formicidae). *Systematic Entomology*, **40**, 61–81
1 Zurita, F., Sánchez, A., Burgos, M., Jiménez, R. and de la Guardia, R.D. (1997).

2 Interchromosomal, intercellular and interindividual variability of NORs studied with

3 silver staining and in *situ* hybridization. *Heredity*, **78**(3), 229-234.