# The innovation of the symbiosome has enhanced the evolutionary stability of nitrogen fixation in legumes

Sergio Miana de Faria, Jens J. Ringelberg, Eduardo Gross, Erik J.M. Koenen, Domingos Cardoso, George K. D. Ametsitsi, John Akomatey, Marta Maluk, Nisha Tak, Hukam S. Gehlot, et al.

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Sergio Miana de Faria, Jens J. Ringelberg, Eduardo Gross, Erik J.M. Koenen, Domingos Cardoso, et al.. The innovation of the symbiosome has enhanced the evolutionary stability of nitrogen fixation in legumes. New Phytologist, 2022, 235 (6), pp.2365-2377. 10.1111/nph.18321. hal-05179640

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# The innovation of the symbiosome has enhanced the evolutionary stability of nitrogen fixation in legumes

Sergio M. de Faria<fnref n="1" /> ID, Jens J. Ringelberg² ID, Eduardo Gross³ ID, Erik J. M. Koenen² ID, Domingos Cardoso⁴ ID, George K. D. Ametsitsi⁵, John Akomatey⁵, Marta Maluk⁶ ID, Nisha Tak⁷ ID, Hukam S. Gehlot⁷ ID, Kathryn M. Wright⁶ ID, Neung Teaumroong⁸ ID, Pongpan Songwattana⁸, Haroldo C. de Lima⁹,¹⁰ ID, Yves Prin¹¹ ID, Charles E. Zartman¹² ID, Janet I. Sprent¹³, Julie Ardley¹⁴ ID, Colin E. Hughes²* ID and Euan K. James⁶* ID

## Summary

* Nitrogen-fixing symbiosis is globally important in ecosystem functioning and agriculture, yet the evolutionary history of nodulation remains the focus of considerable debate. Recent evidence suggesting a single origin of nodulation followed by massive parallel evolutionary losses raises questions about why a few lineages in the N₂-fixing clade retained nodulation and diversified as stable nodulators, while most did not. Within legumes, nodulation is restricted to the two most diverse subfamilies, Papilionoideae and Caesalpinioideae, which show stable retention of nodulation across their core clades.

* We characterize two nodule anatomy types across 128 species in 56 of the 152 genera of the legume subfamily Caesalpinioideae: fixation thread nodules (FTs), where nitrogen-fixing bacteroids are retained within the apoplast in modified infection threads, and symbiosomes, where rhizobia are symplastically internalized in the host cell cytoplasm within membrane-bound symbiosomes (SYMs).

* Using a robust phylogenomic tree based on 997 genes from 147 Caesalpinioideae genera, we show that losses of nodulation are more prevalent in lineages with FTs than those with SYMs.

* We propose that evolution of the symbiosome allows for a more intimate and enduring symbiosis through tighter compartmentalization of their rhizobial microsymbionts, resulting in greater evolutionary stability of nodulation across this species-rich pantropical legume clade.

Authors for correspondence:
Euan K. James
Email: euan.james@hutton.ac.uk

Colin E. Hughes
Email: colin.hughes@systbot.uzh.ch

Received: 3 March 2022

Accepted: 31 May 2022

New Phytologist (2022)

doi: 10.1111/nph.18321

**Key words:** evolution, fixation threads, Leguminosae, nitrogen fixation, nodulation, phylogenomics, symbiosis, symbiosomes.

## Introduction

The N₂-fixing clade of angiosperms includes all plants that form specialized organs known as nodules, within which they house intracellular diazotrophic bacteria (van Velzen et al., 2018a). Within this clade, some species of Cucurbitales, Fagales and Rosales engage in nodulating symbiosis with the filamentous actinobacteria

*Frankia*, while *Parasponia* (Rosales, Cannabaceae) and legumes (Fabales, Fabaceae) host phylogenetically diverse strains of α- and β-proteobacteria collectively known as rhizobia (Soltis *et al.*, 1995; Sprent *et al.*, 2017; Griesmann *et al.*, 2018; van Velzen *et al.*, 2018a). Strikingly, nodulation is mostly a rare trait across these four orders, having been reported in relatively few species except in Fabales, where the majority of the c. 20 000 species in the Fabaceae appear to be nodulated (Doyle, 2011). Across the legume family, nodulation is also very unevenly distributed, with

<footnote n="1">¹Embrapa Agrobiologia, 465 km 07, Seropédica, Rio de Janeiro, BR 23891-000, Brazil; ²Department of Systematic and Evolutionary Botany, University of Zurich, Zollikerstrasse 107, Zurich CH-8008, Switzerland; ³Departamento de Ciências Agrárias e Ambientais, Universidade Estadual de Santa Cruz (UESC), Ilhéus, BA 45662-900, Brazil; ⁴National Institute of Science and Technology in Interdisciplinary and Transdisciplinary Studies in Ecology and Evolution (INCT IN-TREE), Instituto de Biologia, Universidade Federal de Bahia (UFBA), Rua Barão de Jeremoabo, s.n., Ondina, Salvador, 40170-115 BA, Brazil; ⁵CSIR-Forestry Research Institute of Ghana, FUMESUA, PO Box UP 63 KNUST, Kumasi, Ghana; ⁶The James Hutton Institute, Invergowrie, Dundee, DD2 5DA, UK; ⁷Department of Botany, BNF and Microbial Genomics Lab., Center of Advanced Study, Jai Narain Vyas University, Jodhpur, 342001, Rajasthan, India; ⁸School of Biotechnology, Institute of Agricultural Technology, Suranaree University of Technology, Nakhonratchasima 30000, Thailand; ⁹Instituto de Pesquisas Jardim Botânico do Rio de Janeiro (JBRJ/MMA), Rua Pacheco Leão 915, Rio de Janeiro, 22460-030 RJ, Brazil; ¹⁰Instituto Nacional da Mata Atlântica (INMA-MCTI), Av. José Ruschi 4, Santa Teresa, 29650-000 ES, Brazil; ¹¹CIRAD, UMR LSTM, Campus de Baillarguet, 34398, Montpellier Cedex 5, France; ¹²Departamento de Biodiversidade, Instituto Nacional de Pesquisas da Amazônia (INPA), Av. André Araújo Aleixo, Caixa Postal 478, Manaus, 69060-001 AM, Brazil; ¹³Division of Plant Sciences, University of Dundee at The James Hutton Institute, Invergowrie, Dundee, DD2 5DA, UK; ¹⁴College of Science, Health, Engineering and Education, Murdoch University, Murdoch, WA 6150, Australia</footnote>

<footnote>*Joint senior authors.</footnote>
most species in Papilionoideae and the Mimosoid clade (Caesalpinioideae sensu LPWG, 2017) being nodulated, whereas nodulation is less common in nonmimosoid Caesalpinioideae and absent in the other four smaller legume subfamilies (sensu LPWG, 2017). The reasons for this uneven phylogenetic distribution of nodulation are unclear.

Despite the ecological and economic significance of N₂-fixing root nodule symbiosis in ecosystem functioning and agriculture (Peoples et al., 1995; Batterman et al., 2013; Vitousek et al., 2013; Epihov et al., 2017), there is no consensus about the evolutionary origins of this important trait. Hypotheses have shifted from a scenario of multiple origins (Doyle, 2011; Werner et al., 2014), potentially predisposed by a cryptic precursor that evolved in the ancestor of the N₂-fixing clade (Soltis et al., 1995; Werner et al., 2014), to one of a single origin and massive parallel evolutionary losses (Soltis et al., 1995; Griesmann et al., 2018; van Velzen et al., 2018a,b). This second hypothesis was generally dismissed because multiple independent origins provided a more parsimonious solution for the phylogenetic distribution of nodulating lineages, and because variation in nodule types and microsymbionts suggested that nodules are potentially nonhomologous and arose multiple times. This clustered homoplasious occurrence of nodulation, confined to just one clade of angiosperms (Marazzi et al., 2012), prompted the idea that a cryptic precursor evolved in the ancestor of the N₂-fixing clade, which conferred a propensity for nodulation that was expressed in just a subset of lineages (Soltis et al., 1995; Doyle, 2011, 2016; Werner et al., 2014). However, no evidence for such a precursor, genetic or otherwise, has been found (Doyle, 2016; Griesmann et al., 2018; van Velzen et al., 2018a). Furthermore, nodulation involves structural and biochemical innovations underpinned by many genes, multiple developmental and signalling pathways, and coordination between the host and the microsymbiont (Brewin, 2004; Oldroyd & Downie, 2008; Oldroyd, 2013; Sprent et al., 2017; Ardley & Sprent, 2021; Ledermann et al., 2021), such that evolutionary gains of nodulation are likely to be more difficult than losses (van Velzen et al., 2018a; Edwards, 2019). Recently, the alternative hypothesis of a single evolutionary origin of nodulation followed by numerous parallel evolutionary losses has gained traction, notably from comparative genomic studies documenting pseudogenization or loss of key nodulation genes in nonnodulating species, indicative of secondary losses of nodulation (Griesmann et al., 2018; van Velzen et al., 2018a; Zhao et al., 2021). Reexamination of the structural and developmental homologies and commonalities in symbiotic gene function across nodulating lineages spanning the N₂-fixing clade suggested that these also provide more compelling evidence for the single gain and multiple losses hypothesis (van Velzen et al., 2018a; Shen & Bisseling, 2020).

This shift in thinking prompts questions about how the numerous secondary losses of nodulation are distributed across lineages and through time, why certain lineages retained nodulation to diversify as stable N₂-fixers whereas many others lost this trait, and why, through time, N₂-fixing symbiosis apparently became nonadvantageous for the large majority of N₂-fixing clade lineages.

One trait that has not been considered as a potential determinant of evolutionary stability of nodulation is the occurrence of two distinct anatomical arrangements of N₂-fixing bacteria within the nodule. In the majority of papilionoid legumes, such as pea and *Medicago*, an infection thread (IT), formed from invagination of a root hair cell wall, conveys rhizobia from the point of infection to the nodule primordium. Rhizobia within the IT are budded off once they reach the nodule cell and are retained within it only by the host plasmalemma-derived symbiosome (or peribacteroid) membrane, where they differentiate into their N₂-fixing bacteroid forms (Sprent, 2001, 2009; Brewin, 2004; Sprent et al., 2017; Parniske, 2018; Ardley & Sprent, 2021; Tsyganova et al., 2021). By contrast, in all actinorhizal symbioses and in a subset of nodulating legumes the N₂-fixing bacteria are retained within modified, thin-walled, infection threads called fixation threads (FTs), remaining enclosed within the plant cell wall and the plasmalemma. Hereafter, we refer to these as FT-type nodules, and those in which the bacteroids are enclosed in symbiosomes as SYM-type nodules.

FT-type nodules were first described in actinorhizal plants (enclosing their *Frankia* microsymbionts), and in *Parasponia*, the only nonlegume known to form nodules with rhizobia (Trinick, 1980; Lancelle & Torrey, 1985; Smith et al., 1986). They were later observed in legumes, mostly in woody Caesalpinioideae (*sensu* LPWG, 2017) where they appeared to be relatively common (de Faria et al., 1986, 1987; Naisbitt et al., 1992; Sprent, 2001; Fonseca et al., 2012). FT-type nodules have also been reported in a few legumes belonging to the subfamily Papilionoideae, which is sister to Caesalpinioideae (Koenen et al., 2020b; Zhao et al., 2021), including tree genera such as *An dira*, *Dahlstedtia* and *Hymenolobium*, and members of tribe Brongniartieae (de Faria et al., 1986, 1987; Sprent, 2001, 2009; Sprent et al., 2013, 2017). Ultrastructural and histochemical analyses of FT-type nodules in *Parasponia* with rhizobia (Smith et al., 1986), actinorhizal nodules with *Frankia* (Pawlowski & Demchenko, 2012) and in some legumes (de Faria et al., 1986, 1987; Naisbitt et al., 1992) revealed that FTs are superficially similar to the cell wall-bound ‘invasive’ IT, for example in harbouring some pectin (Fonseca et al., 2012). The IT is an extension of the host cell wall and comprises mainly cellulose and pectin; its role appears to be largely protective, preventing the bacteria from invading the plant in a disorganized or pathogenic manner (Brewin, 2004; Tsyganova et al., 2021). However, the composition and role of the FT remain uncertain. Moreover, the precise nature of the FT in relation to the symbiosome membrane, which in SYM-type nodules is essential for the exchange of nutrients between the host cytoplasm and the bacteroid (White et al., 2007), remains unknown.

Within legumes, nodulation is restricted to the two largest subfamilies, Caesalpinioideae (which includes the phylogenetically nested Mimosoid clade, formerly subfamily Mimosoideae – see LPWG, 2017) and Papilionoideae (Sprent et al., 2017; Ardley & Sprent, 2021). Here we investigate the occurrence of FT- and SYM-type nodulation across Caesalpinioideae, the second largest subfamily of legumes, with 152 genera and c. 4600 species
distributed pantropically across all lowland tropical biomes, with minor incursions into temperate regions. We provide an updated census of nodulation occurrence, including three new records in genera of previously unknown nodulation status, plus extensive new data regarding FT- and SYM-type nodules across genera. We investigate whether these two nodule types represent different degrees of 'compartmentalization' (i.e. physical structures that allow hosts to spatially separate symbionts; Chomicki *et al.*, 2020), by examining the anatomy and structure of nodules in *Chidlowia*, *Pentaclethra* and *Erythrophleum*, genera that span an evolutionary transition from FT- to SYM-type nodules that we hypothesize to have occurred within Caesalpinioideae along the branch subtending the Mimosoid clade (Fig. 1). *Erythrophleum*, which is placed in the sister clade of the Mimosoid clade, has FT-type nodules, while *Pentaclethra* and *Chidlowia*, which are among the first-branching lineages of the Mimosoid clade (Manzanilla & Bruneau, 2012; Koenen *et al.*, 2020a), as well as all other studied taxa in the Mimosoid clade, have SYM-type nodules.

We test the hypothesis that the transition in nodule anatomy from FT- to SYM-type nodules constituted an evolutionary innovation that led to more stable retention of nodulation, whereas lineages in which FT-type nodules occur are more prone to evolutionary losses of nodulation. For this, we explore the number and phylogenetic distribution of evolutionary losses of nodulation using a robust phylogenomic backbone that includes 97% of Caesalpinioideae genera. We also examined the composition of the FT wall in more detail than has hitherto been achieved using

immunohistochemical methods that have been used for ITS (Brewin, 2004; Tsyganova et al., 2021) in order to better elucidate its possible role in symbiosis.

**Fig. 1** Evolutionary trajectories of nodulation and nodule type across a time-calibrated phylogeny of the legume subfamily Caesalpinioideae. Pie charts on nodes show the proportions of the most probable reconstructed character states: nonnodulating, fixation thread (FT-type nodules), symbiosome (SYM-type nodules), nodulating but of unknown type, and nodulation status unknown, summarized over 500 simulations. Branch colours denote the nodulation status of the node or tip it subtends and the coloured boxes in front of each taxon name show the character state for that species. Note that in three clades (the *Senna* + *Cassia* clade, the *Arcoa* + *Acrocarpus* clade and the *Peltophorum* clade) 'double' losses of FT-type nodules are inferred to have occurred simultaneously in both descendant lineages of that node. For example, the crown node of the *Senna* + *Cassia* clade is inferred to be nodulating with FT-type nodules even though *Senna* and *Cassia* are nonnodulating. The dashed orange, blue and dark green vertical lines show the phylogenetic locations and maximum ages of the various character state transitions on the tree. Using the same colours, the histograms show the frequencies of the number of transitions from FT to SYM (blue), from SYM to nonnodulation (green) and from FT to nonnodulation (orange), and the rates of losses of nodulation per million years for SYM to nonnodulation (green) and FT to nonnodulation (orange) across 500 independent character estimations. Note that while the three other character state transitions, from nonnodulating to FT or SYM-type nodules, and from SYM to FT, were not allowed under our model, and were therefore fixed at zero, an alternative model permitting all character state transitions gave an identical result (Supporting Information Fig. S3). Asterisks after terminal names indicate species which are the focus of detailed nodule anatomical work presented in Figs 2–4. Ple, Pleistocene; Pli, Pliocene.

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Materials and Methods

Nodulation and nodule anatomy

Basic nodulation data (nodulated or nonnodulated) were
obtained from Sprent (2001, 2009), from papers or reports pub-
lished since 2009, and from previously unpublished records in
the databases of the authors, including new reports of nodulation
status (Supporting Information Tables S1 and S2). Where no
data are available the nodulation status of a genus is listed as
Uncertain (Un) (Table S1).

Anatomical types (FT or SYM) were determined from exten-
sive data newly obtained here (Tables S1 and S2) alongside pub-
lished data for the specific taxa that were used to construct the
phylogeny, or related species in the same genus, based on sub-
stantial data that indicate that nodulation is almost always a
generic trait. However, losses of nodulation have been reported
within the genus *Senegalia* (see below), and it should be acknowl-
edged that the relatively limited nodulation record and sampling
in the Caesalpinioideae may hide additional intrageneric losses
(Sprent *et al.*, 2017). All samples were prepared for light and elec-
tron microscopy according to de Faria *et al.* (1986, 1987) and
Fonseca *et al.* (2012), unless otherwise stated in Table S2; in total,
samples from 95 species were newly prepared for this study.
Additional samples of nodules from *Chidlowia sanguinea*, *Entada
polystachya*, *Erythrophleum* spp., *Moldenhawera* spp. and *Penta-
clethra macroloba* were prepared specifically to examine in detail
the presence of FTs or symbiosomes (Table S2, Notes S1). For
these samples, slices from four or more nodules per species were
fixed in 2.5% glutaraldehyde and processed in two ways: (1) for
light microscopy and immunogold transmission electron
microscopy (TEM) with the monoclonal antibodies JIM5 and
JIM7, which recognize, respectively, unesterified and esterified
pectin (VandenBosch *et al.*, 1989; Tsyganova *et al.*, 2021), accord-
ing to Fonseca *et al.* (2012); and (2) for identifying the symbio-
some membrane by TEM using additional postfixation in osmium
tetroxide followed by embedding in epoxy resin according to
Rubio *et al.* (2009). Ultramicrotomy, staining of sections for light
microscopy and for TEM, and immunogold labelling with JIM5
for TEM were as described in Fonseca *et al.* (2012).

For immunohistochemical analysis of the FT wall, confocal
laser scanning microscopy (CLSM) was performed on slides con-
taining semithin sections (1 µm thickness) of *Erythrophleum* and
*Pentaclethra* nodules fixed and embedded as per method (1)
above. The sections were incubated for 2 h in 1 : 10 dilutions of
monoclonal antibodies raised against various plant cell wall com-
ponents (all obtained from Plant Probes, Centre for Plant
Sciences, University of Leeds, UK): Lm2, which labels β-linked-
GlcA in arabinogalactose protein (AGP) glycan; Lm5, which
labels the pectic polysaccharide rhamnogalacturonan; and Lm15,
which labels the XXXG motif of the nonpectic, noncellulosic
polysaccharide xyloglucan. Then, after washing twice in distilled
water (dH₂O) the sections were incubated for 1 h in a 1 : 500
dilution of goat antirat Alexa 488 secondary antibody
(ThermoFisher, Loughborough, UK) followed by several rinses
with dH₂O. After mounting in coverslips and Fluoromount

(ThermoFisher), the sections were examined using a Zeiss LSM
710 confocal laser scanning microscope (Carl Zeiss Microscopy
Ltd, Cambourne, UK), fitted with a W Plan-Apochromat 40×
lens, using spectral imaging with excitation at 488 nm and emis-
sions between 494 nm and 727 nm. The images were colour-
coded according to wavelength and enhanced using the Min/
Max function in ZEN 2010 software. Ultrathin sections (80 nm)
of the same samples were then immunogold labelled for TEM
using the same monoclonal antibodies as those for CLSM (Lm2,
Lm5, Lm15) according to Fonseca *et al.* (2012).

Phylogeny and ancestral trait estimation

We used a recently constructed time-calibrated phylogeny of
Caesalpinioideae that included 147 of the 152 genera, which was
based on DNA sequence data derived from targeted enrichment
of 997 nuclear genes selected specifically for phylogenomics of
the Mimosoid clade using the Mimobaits bait set (Koenen
et al., 2020a) to generate a large phylogenomic Hybseq dataset
(Ringelberg et al., 2022). Using this much larger gene set allowed
us to overcome lack of resolution prevalent across the backbone
of the nonmimosoid grade in previous phylogenies that were
based on much smaller traditional Sanger DNA sequence datasets
(Brunneau et al., 2008; Manzanilla & Bruneau, 2012; LPWG,
2017) to generate a robust and densely sampled phylogenetic
hypothesis. The five unsampled genera are *Hultholia*, a member
of the Caesalpinia clade (Gagnon et al., 2016), a group of 27 gen-
era that are all either nonnodulating or of unknown nodulation
status; *Stenodrepanum*, which is sister to *Hoffmannseggia*, and
is also placed in the Caesalpinia clade (Gagnon et al., 2016);
*Pterogyne*, which is also nonnodulating and probably forms a
phylogenetically isolated monogeneric lineage in the nonmi-
mosoid grade of Caesalpinioideae that is potentially sister to a
large clade comprising all Caesalpinioideae except the Umtiza
and Ceratonia clades (Zhao et al., 2021); *Microlobius*, which is
nodulating with SYM-type nodules and probably nested within
the genus *Stryphnodendron* (Simon et al., 2016; Ribeiro et al.,
2018); and finally the nonmimosoid *Vouacapoua*, which is
nonnodulating, but is probably placed in the Cassia clade,
which contains both nodulating and nonnodulating lineages
(Brunneau et al., 2008).

The original 420-taxon Caesalpinioideae phylogeny of Ringel-
berg et al. (2022) was time-calibrated in BEAST (Drummond &
Rambaut, 2007), using a species tree topology estimated by
ASTRAL (Zhang et al., 2018), a subset of 100 informative and
clock-like genes, and seven Caesalpinioideae fossil constraints
(Ringelberg et al., 2022). We pruned this chronogram until each
genus was represented by just a single taxon, with two exceptions:
first, the genus *Chamaecrista*, for which we retained four species
due to known variation in nodule type within that genus (Nais-
bitt et al., 1992; Santos et al., 2017); and second, the dense sam-
pling of 420 taxa in the original phylogeny allowed us to test the
monophyly of genera, and for nonmonophyletic genera we
retained representative taxa for each para-/polyphyletic lineage.
Nodulation data (Table S1) were matched to the tips of this tree
in as conservative a way as possible. For example, in the case of
*Prosopis*, which is polyphyletic and is thus represented by four taxa in the tree, these were scored as follows: *P. juliflora*, *P. cineraria* and *P. africana*, which represent three independent lineages, are scored as SYM, as the nodule type of these taxa is known. By contrast, the taxon representing the fourth lineage, *P. ferox*, is scored as nodulating but with an unknown nodule type, because *P. ferox* and other taxa in this clade are nodulating, but their nodule types are unknown. In the case of *Senegalia*, which is also nonmonophyletic and where losses of nodulation have been reported (Sprent, 2001, 2009), but where the phylogenetic distribution of those losses remains unknown, we ran an additional trait reconstruction including two losses within the genus. The extensive generic nonmonophyly visible in *Caesalpinioideae*, especially within the mimosoid clade (Fig. 1), is the focus of current taxonomic work reported elsewhere (J. J. Ringelberg et al., unpublished).

Nodulation status was estimated across this phylogeny using a model with three character states: nonnodulating, fixation thread (FT), and symbiosome (SYM). We explicitly followed the single gain, multiple losses evolutionary model for the origins of nodulation; that is, nodulation evolved only once in angiosperms and was subsequently lost many times. To do this, we constrained the model so that the root state of the tree was set to FT and ran the reconstructions either allowing only three types of transitions: from FT to nonnodulating, from FT to SYM, and from SYM to nonnodulating, or allowing all possible transitions. We used stochastic character mapping implemented in the function make.simmap in the PHYTOOLS R package (Revell, 2012) to simulate 500 independent evolutionary trajectories of nodulation across the time-calibrated phylogeny. Results were summarized onto this tree across all simulations, with transitions inferred along branches connecting nodes that have different character states in the majority of the 500 simulations. Rates of character change (i.e., events per million years) were extracted from the transition matrix of the continuous-time-reversible Markov model fitted to the data by the make.simmap function. Taxa for which the nodulation status is unknown were assigned equal probabilities for all three character states.

## Results

### Nodulation and nodule anatomy

Data on nodule anatomy for 128 species of 56 genera of *Caesalpinioideae*, including more than 80 records newly reported here (Tables S1, S2), show that no species belonging to the Mimosoid clade have FT-type nodules; that is, all known nodulating mimosoids have SYM-type nodules (Figs 1, S1). By contrast, all nodulating species from the grade subtending the Mimosoid clade have FT-type nodules (Figs 1, S2), except for a subset of species of *Chamaecrista*. In that sense *Chamaecrista* is exceptional among the taxa of the nonmimosoid grade, as it harbours species with either FT- or SYM-type nodules (Naisbitt et al., 1992).

Mature nodules of *Erythrophleum ivorense* and *E. suaveolens*, placed in the sister group of the Mimosoid clade (Fig. 1), are of

the indeterminate type (Sprent, 2001, 2009; Sprent et al., 2017), and similar to those described in detail in the genus *Dimorphandra*, which is related to *Erythrophleum* (Fonseca et al., 2012). Accordingly, as with *Dimorphandra*, nodules have a meristem at the tip and a large invasion zone (IZ) (Fig. 2a), containing cells being invaded by rhizobia (Fig. 2b), and an N₂-fixing zone occupying most of the nodule volume; this contains a mix of infected and uninfected cells (Fig. 2a). Bacteria can be seen to invade the IZ cells via ITs emerging from between the cells (Fig. 2c); these ITs are intensely immunogold-labelled with the monoclonal antibody JIM5, indicating that they contain unesterified pectin, specifically partially methylated homogalacturonan (VandenBosch et al., 1989; Tsyganova et al., 2021), and are similar in that respect to the host cell wall (Fig. 2c,d). Similar results were obtained with JIM7 (data not shown), indicating that *Erythrophleum* ITs also contain esterified pectin in common with papilionoid nodules (Tsyganova et al., 2021). Infected cells in the N₂-fixing zone contain numerous FTs (Fig. 2e), bound by cell walls that are thinner than those of the ITs, and either have sparser labelling with JIM5 (Fig. 2d,f) and JIM7 (not shown) or have no apparent labelling (Fig. 2f). Higher definition TEM with osmicated samples reveals that the FTs are surrounded by a cell membrane (Fig. 2g,h), which appears to be derived from the host endoplasmic reticulum (ER). This shows that the FT comprises a multilayered compartment consisting of a cell membrane, the FT cell wall, the lumen of the FT and the bacteroid (Fig. 2g). Infected cells of nodules of the neotropical legumes *Moldenhawera floribunda* and *M. blanchetiana* var. *multijuga*, also placed in the nonmimosoid grade of caesalpinioids (Fig. 1), are packed with bacteroids enclosed within FTs (Fig. S2a,b). As with *Erythrophleum*, the ITs are intensely labelled with JIM5 (Fig. S2c), but the FTs considerably less so (Fig. S2d); the FTs in *Moldenhawera* are also associated with membranes arising from the ER (Fig. S2e,f). Another three nonmimosoid neotropical caesalpinoid genera (*Jacqueshuberia purpurea*, Fig. S2g,h; *Tachigali rugosa*, Fig. S2i,j; and *Campsiandra comosa*, Fig. S2k,l) also have their bacteroids enclosed in FTs labelled to various degrees with JIM5. Together with published reports on *Chamaecrista* (de Faria et al., 1987; Naisbitt et al., 1992) and *Dimorphandra* (Fonseca et al., 2012), these observations of five additional genera demonstrate the ubiquity of FT-type nodules across nodulating lineages in the nonmimosoid grade of *Caesalpinioideae* (Fig. 1).

By contrast, *Pentaclethra macroloba*, which is placed among the first-branching lineages of the mimosoid clade (Fig. 1), has indeterminate nodules (Fig. 3a) with infected cells in the N₂-fixing zone surrounded by uninfected cells (Fig. 3b) containing bacteroids that are not surrounded by a cell wall (Fig. 3c) but are clearly enclosed within symbiosomes (Fig. 3d). In the same clade, *Xyla xylocarpa* also has SYM-type nodules (Fig. S1g,h). *Chidlowia sanguinea* nodules are similar to those of *P. macroloba*, except that the IZ is more prominent (Fig. 3e); the bacteroids are also enclosed in symbiosomes (Fig. 3f). *Chidlowia sanguinea* is sister to a large clade containing the bulk of mimosoid species, wherein SYM-type nodules are consistently present, as illustrated by *Entada polystachya* (Fig. S1a,b), *Enterolobium cyclocarpum*
<figure><img src="image_5.png" /><figcaption>**Fig. 2** Nonmimosoid grade caesalpinioid nodules in the genus *Erythrophleum* contain bacteroids enclosed within fixation threads (FTs). Light (a, b) and transmission electron microscope (c–h) images of sections of nodules from *E. ivorense* (a–f) and *E. suaveolens* (g, h). (a) Whole nodule longitudinal profile illustrating the zonation typical of an indeterminate nodule (m, meristem; iz, invasion zone; nf, nitrogen fixing zone). Bar, 250 µm. (b) Higher magnification view of the iz in which newly divided host cells derived from the meristem (m) are being invaded by numerous infection threads (arrows). Bar, 10 µm. (c) Large infection thread containing bacteria (b) invading cells in the iz; the walls of the infection thread are densely immunogold labelled with 10 nm gold particles linked to JIM5 (arrows), a monoclonal antibody which recognizes nonesterified pectin. v, vacuole. Bar, 1 µm. (d) Infection thread in the iz-nf boundary with its cell walls labelled with JIM5 (large arrows) adjacent to an FT (*) with a thinner cell wall that is very sparsely labelled with JIM5 (arrowheads). Bar, 500 nm. (e) Cell in the nf zone packed with FTs (black arrows), including within the nucleus (n) indicated by white arrowheads. Bar, 2 µm. (f) Detail of FTs in the nf zone containing N-fixing bacteroids (b); the FT walls range from being sparsely labelled with JIM5 (arrowheads) to exhibiting little or no obvious labelling (single gold particles are indicated by arrows). Bar, 500 nm. (g) High-resolution image of a bacteroid (b) forming within a strand of cytoplasm between vacuoles (v) in an iz cell; the bacteroid is surrounded by a cell wall (w) that is being enveloped in a membrane (arrowheads), stretches of which (arrows) appear to be derived from nearby endoplasmic reticulum/Golgi bodies (er). The intense metabolic activity of this process is suggested by the nearby mitochondria (m) and peroxisomes (p). Bar, 1 µm. (h) Bacteroids (b) in newly formed FTs packed into a new N-fixing cell in the early nf zone adjacent to the iz; the bacteroids are surrounded by the FT wall (w), which is itself surmounted by a symbiosome membrane (arrowheads). Note the membranes within the cytoplasm that are associated with the FTs (arrows). n, nucleus; m, mitochondrion. Bar, 1 µm.</figcaption></figure>

(Fig. S1c,d) and *Lachesiodendron viridiflorum* (Fig. S1e,f), showing that symbiosomes are universally found in nodulating lineages across the entire Mimosoid clade (Fig. 1; Table S1).

The FT wall in *Erythrophleum* nodules was investigated further using monoclonal antibodies against AGP glycan (Lm2), the pec- tic polysaccharide rhamnogalacturonan (Lm5) and the nonpectic,
**Fig. 3** Nodules of caesalpinioids from the Mimosoid clade contain bacteroids enclosed within symbiosomes (SYMs). Light (a, b, e) and transmission electron microscope (c, d, f) images of sections of nodules from *Pentaclethra macroloba* (a–d) and *Chidlowia sanguinea* (e, f). (a) Whole *P. macroloba* nodule longitudinal profile illustrating the zonation typical of an indeterminate nodule (m, meristem; nf, nitrogen fixing zone). Bar, 500 µm. (b) Higher magnification view of the nf zone showing large bacteroid-containing cells (b) surrounded by smaller and more numerous uninfected cells (u). n, nucleus. Bar, 25 µm. (c) Bacteroids (b) within a symbiosome adjacent to the host cell wall (w) which is immunogold labelled with 10 nm gold particles linked to JIM5 (arrows). The symbiosome peribacteroid space is marked with an asterisk (*); note that there is no cell wall separating the symbiosome from the host cytoplasm (c). Bar, 500 nm. (d) High-resolution image of bacteroids (b) housed in symbiosomes; the symbiosome membrane separating it from the cytoplasm (c) are marked with arrows, and the peribacteroid space by an asterisk (*). m, mitochondrion. Bar, 500 nm. (e) Whole *C. sanguinea* nodule longitudinal profile illustrating the zonation typical of an indeterminate nodule (m, meristem; iz, invasion zone; nf, nitrogen fixing zone). Bar, 200 µm. (f) High-resolution image of bacteroids (b) housed in symbiosomes; the symbiosome membrane separating it from the cytoplasm (c) is marked with arrows, and the peribacteroid space by an asterisk (*). Bar, 500 nm.

noncellulosic polysaccharide xyloglucan (Lm15). These probes were capable of clearly delineating FTs by both CLSM (Fig. 4a–d) and TEM (Fig. 4e–h), indicating that the walls of FTs contain all three of these components. By contrast, the symbiosomes in *Pentaclethra* nodule sections treated identically were very difficult to discern using CLSM (Fig. 4j–l), and although they could be observed under TEM they had few or no gold particles associated with them, indicating an absence of cell wall components (Fig. 4n–p). The exception was Lm2 (AGP glycan), which labelled *Pentaclethra* symbiosomes (Fig. 4i,m), and thus confirmed previous observations of AGP in the symbiosome membrane made with pea (*Pisum sativum* L.) nodules (Tsyganova et al., 2021).

Phylogeny and evolution of nodule types

Ancestral estimation of nodulation and nodule types across the caesalpinioid phylogeny reveals two independent transitions from FT- to SYM-type nodules, first on the branch subtending the Mimosoid clade in the mid-Eocene, 46–41 million years ago (Ma) (95% highest posterior density (HPD): 46.5–40.7 Ma),

and later within the genus *Chamaecrista* (P. A. Casaes et al., unpublished) in the early to mid-Miocene, 27–13 Ma (95% HPD: 27.6–12.4 Ma) (Fig. 1), or potentially later given our sparse sampling of species of *Chamaecrista*. Seventeen losses of nodulation are hypothesized across Caesalpinioideae, 12 in ancestrally FT-type lineages and five in ancestrally SYM-type lineages (Fig. 1; Table S3), implying rates of losses of 0.0174 per million years (ancestrally FT-type) and 0.0028 (ancestrally SYM-type), that is up to six times greater for FT-type lineages than SYM-type lineages (Fig. 1; Table S4), suggesting that nodulation based on FT-type nodules is significantly more prone to loss than nodulation based on SYM-type nodules. Additional analyses show that allowing all transitions between character states results in an identical reconstruction of losses (Fig. S3). Alternative scoring of character states for taxa with missing data (e.g. assigning equal weight to nodulation and nonnodulation states), or inclusion of losses within *Senegalia* do not significantly affect the outcome (Figs S4, S5). All this suggests that the data strongly support our model of evolution of nodule types across Caesalpinioideae and the higher rate of losses associated with FT-type lineages (Fig. 1).

<figure><img src="image_6.png" /></figure>
<figure><img src="image_7.png" /><figcaption>**Fig. 4** Fixation threads (FTs) contain cell wall components in addition to unesterified (JIM5) and esterified (JIM7) pectin. Confocal laser scanning microscopy (CLSM) with anti-rat Alexa Fluor 488 (a–d, i–l) and immunogold transmission electron microscopy with anti-rat 10 nm gold (e–h, m–p) of *Erythrophleum* (a–h) and *Pentaclethra* (i–p) nodules incubated in monoclonal antibodies raised in rat against various plant cell wall components: Lm2 (a, e, i, m), which labels arabinogalactose protein (AGP) glycan; Lm5 (b, f, j, n), which labels the pectic polysaccharide rhamnogalacturonan; and Lm15 (c, g, k, o), which labels the XXXG motif of the nonpectic, noncellulosic polysaccharide xyloglucan. Control sections incubated in buffer alone without a primary antibody are presented in (d, h, i, p). FTs are indicated by arrows in (a–h), and symbiosomes by arrowheads in (i–p). w, host cell wall separating plant cells; b, bacteroid. Bars: (a–d, i–l) 5 µm; (e–h, m–p) 1 µm.</figcaption></figure>

Maximum ages of evolutionary losses of nodulation from FT-
type nodule ancestry span the late Palaeocene to the late Oligocene
(59–24 Ma (95% HPD: 59.2–23.0 Ma)) and from SYM-type nod-
ule ancestry from the late Eocene to the late Oligocene (37–25 Ma
(95% HPD: 37.2–24.6 Ma)) (Fig. 1; Table S3).

Discussion

Within legumes, nodulation is restricted to the two largest sub-
families, Caesalpinioideae (sensu LPWG, 2017) and Papil-
ionoideae. The idea that nodulation was ‘stabilized’ in these two
lineages was noted by Werner *et al.* (2014), who referred to core clades of Papilionoideae as 'stable fixers' (i.e., a clade across which losses of nodulation were almost absent) and to the Mimosoid clade (subfamily Mimosoideae in Werner *et al.*, 2014) as having a 'moderately stable fixing state', where losses of nodulation were infrequent. Here we provide an explanation for this pattern: that the evolution of SYM-type nodules, with bacteroids contained within a symbiosome, accounts for this greater stability. FT-type nodules characterize almost all nodulating nonmimosoid Caesalpinioideae genera, while the transition to SYM-type nodules on the stem lineage of the Mimosoid clade (Fig. 1) coincides with a shift to fewer losses of nodulation, and to a significantly lower rate of losses of nodulation per million years (i.e. a shift to greater evolutionary stability of N₂-fixation in this clade). As the FT-type nonmimosoid Caesalpinioideae lineages are relatively more densely sampled in our phylogeny (Fig. 1) than the SYM-type mimosoids, the difference between the rates of loss of FT and SYM is probably underestimated; the difference in rates of losses from FT-type lineages compared to SYM-type lineages is likely to be even greater than we report here.

In Papilionoideae, the sister group of Caesalpinioideae (LPWG, 2017; Koenen et al., 2020b), nodulating and nonnodulating genera appear to be similarly concentrated phylogenetically across the initial divergences of the subfamily, while later stabilizing as nodulating in most lineages within the large 50 kb inversion clade (Werner et al., 2014; Doyle, 2016; Epihov et al., 2017; van Velzen et al., 2018a). It remains to be tested whether there is a similar association between more frequent losses of nodulation and FT-type nodules in Papilionoideae, where FT-type nodules are also found sporadically, but the vast majority of nodulating lineages and all the species-rich nodulating clades – the ‘stable fixers’ (Werner et al., 2014) – have SYM-type nodules. Lack of phylogenetic resolution among the initial divergences in Papilionoideae (Cardoso et al., 2012, 2013; LPWG, 2017) means that this test must await a more robust phylogeny.

Assuming a single gain of nodulation followed by massive losses, the consistent occurrence of FT-type nodules across nodulating lineages in the nonmimosoid grade of Caesalpinioideae demonstrates that FT-type nodules are ancestral within Caesalpinioideae and persisted through early evolution of that subfamily (Fig. 1). The FT-type nodule also characterizes actinorhizal nodules (Pawlowski & Demchenko, 2012) and rhizobial nodules in *Parasponia* (Lancelle & Torrey, 1985) and is thus most likely to be ancestral across the N₂-fixing clade (Shen & Bisseling, 2020) and legumes as a whole. As noted by Koenen et al. (2020b), rapid initial divergence of the six legume subfamilies implies additional losses of nodulation along the stem lineages or early in the crown group divergences of subfamilies Cercidoideae, Detarioideae, Dialioideae and Duparquetioideae, or potentially even more numerous parallel losses in more recent times, as documented within the genus *Trema* (Cannabaceae) (van Velzen et al., 2018a), as no extant members of these subfamilies are known to nodulate. Although we have not looked at actinorhizal symbioses here, the very low proportion of nodulated species within Cucurbitales, Fagales and Rosales (Ardley & Sprent, 2021) suggests that many lineages have suffered losses of

nodulation similar to the Cercidoideae, Detarioideae, Dialioideae and Duparquetioideae, and the nonmimosoid Caesalpinioideae. However, although they might also be considered as housing their symbionts within FTs, the filamentous nature of *Frankia* precludes it from forming symbiosomes. Furthermore, the ability of *Frankia* to fix N₂ *ex planta* and regulate gaseous and nutrient exchange via vesicles in nodules (*Casuarina* and *Allocasuarina* in the Fagales being the exceptions in that *Frankia* does not form vesicles in the nodules, and the host regulates gaseous exchange) means that *Frankia* has a fundamentally different relationship with its host compared to that between rhizobia and legumes (Pawlowski & Demchenko, 2012; Ardley & Sprent, 2021). Although losses of nodulation amongst actinorhizal lineages are the result of evolutionary drivers that are not currently well understood, it is possible that intimacy (or lack thereof) in terms of the interface between the two symbionts is not an important factor.

It is well established that N₂-fixation is energy-demanding, limited by photosynthesis, and confers fitness advantages only when nitrogen is limiting and when the benefits derived from greater availability of nitrogen (e.g. in fostering higher photosynthetic rates in N₂-fixing plants) are greater than the costs of photosynthetic carbon (McKey, 1994; Hoffman et al., 2014; Taylor & Menge, 2018; van Velzen et al., 2018a). Additionally, there is experimental evidence showing that legumes increase N₂-fixation at elevated CO₂ levels and that nitrogenase activity declines rapidly above 35°C and below 25°C (Trinick, 1980). Taken together, this suggests a greater advantage in being an N₂-fixer under early Cenozoic CO₂ levels and temperatures (Rogers et al., 2009; Chen & Markham, 2021), and for those advantages to be preferentially retained in the tropics and subtropics, where FT-type nodulators are largely restricted. Falling atmospheric CO₂ levels and temperatures through the Cenozoic could have triggered global evolutionary losses of nodulation across the N₂-fixing clade (as suggested by van Velzen et al., 2018a) and we show that maximum ages of losses of nodulation across Caesalpinioideae are widely scattered from the late Palaeocene to the late Oligocene, 59–24 Ma (Fig. 1). However, it is important to consider that these are maximum ages rather than precise indicators of the timing of losses, and furthermore, the number of estimated losses per lineage is the minimum number of losses to explain the observed pattern at the tips, given the taxa sampled here.

It has long been recognized that evolutionary conflicts arise between hosts and symbionts over symbiont mixing, proliferation and transmission (Frank, 1996) because the presence of multiple, genetically heterogeneous symbiont strains within a host will cause symbionts to evolve traits that increase symbiont proliferation, competition and conflict, but decrease the efficiency of the symbiosis (Frank, 1996). There is clear evidence that this happens in the rhizobia–legume symbiosis (Oono et al., 2009; Sachs et al., 2018). In response to such conflicts, hosts have evolved ways to control symbiont proliferation (i.e. terminally differentiated bacteroids lose their ability to reproduce) (Mergaert et al., 2006; De La Peña et al., 2018; Ardley & Sprent, 2021), discriminate among symbionts (Yang et al., 2017; De La Peña et al., 2018; Ardley & Sprent, 2021) and penalize noncooperating symbionts (Kiers et al., 2003; Oono et al., 2009; Ardley & Sprent, 2021). All these approaches are more
effective when symbionts are compartmentalized within hosts, and
all occur in the rhizobia–legume symbiosis (Sachs *et al.*, 2018),
although almost all evidence is from SYM-type papilionoids. We
suggest that the anatomical differences between legume FTs and
SYMs represent different degrees to which symbionts are effec-
tively compartmentalized, as argued by Chomicki *et al*. (2020). The
SYM-type nodule, in which the microsymbiont is released from the
wall-bound IT into membrane-bound symbiosomes within the host
cell, allows for a more intimate and potentially more effective and
enduring symbiotic partnership where the plant has invested in the
establishment of an N₂-fixing ‘organelle’, which Parniske (2018)
argued has only occurred in legumes and in the *Gunnera–Nostoc*
symbiosis. In SYM-type nodules, the plant host assumes greater
control of the microsymbiont and supplies more of the components
required for bacteroid metabolism and N₂-fixation (Hakoyama
et al., 2009; Udvardi & Poole, 2013). This more intimate endosym-
biosis reaches its pinnacle in the Inverted Repeat Lacking Clade
(IRLC) of Papilionoideae, in which swollen endoreduplicated bac-
teroids that have lost their capacity for free-living growth, but which
are highly efficient at fixing N, are prevalent (Oono et al., 2009;
Ardley & Sprent, 2021). The bacteroid in the FT, although sur-
rounded by a membrane analogous to the SYM membrane, remains
surrounded by a cell wall, albeit a thin one which contains little
pectin, which means that it is extracellular (i.e. in the apoplast).
Thus, while FTs represent a modest degree of compartmentaliza-
tion, at least in *Parasponia* there is evidence to suggest that FTs are
not effective in controlling growth of inefficient rhizobial strains
(Op den Camp et al., 2012). These differing degrees of compart-
mentalization provide a compelling reason why SYM-type nodula-
tors, especially those in the IRLC (Westhoek et al., 2021), are less
likely to be affected by cheating or infiltration by inefficient
microsymbionts compared to FT-type nodulator hosts, which
remain in a ‘looser’ relationship with their symbionts.

It might be assumed that the wall of the FT would also present
an additional barrier in terms of nodule O₂ relationships and host-
symbiont nutrient exchange. However, the FT wall is thin, consist-
ing mainly of cellulose, hemicelluloses and AGP, but with reduced
levels of homogalacturonan (HG) pectin components compared to
the thicker and stiffer IT wall (this study), and hence presumably
relatively permeable to C4-dicarboxylates and ammonia
(Brewin, 2004). The presence of abundant leghaemoglobin (Lb)
suggests that rhizobial FT nodules are not obviously different from
SYM-type nodules in their O₂ exchange. We propose that the role
of the thin-walled FT appears to be protective; that is, it prevents
the undifferentiated bacteroids going ‘rogue’ and proliferating
within the host cell, but in parallel, prevents the plant from identi-
fying them as pathogens and attacking them. In all other respects,
FT-type nodules are similar to SYM-types in still possessing a sym-
biosome membrane which surrounds the FT and is the real inter-
face of nutrient exchange between the two partners. In short, the
FT represents an example of a symbiosis that functions well
enough to benefit both partners, but is a looser, more metaboli-
cally independent association.

SYM-type nodules may accommodate a larger diversity of
symbionts and facilitate adaptation to temperate climates. This is
in line with the greater diversity of α- and β-rhizobial types in

SYM-type nodules and the wider geographical distribution and
environmental span of SYM-type nodulating legumes (Sprent
et al., 2017; Ardley & Sprent, 2021), compared to FT-type nod-
ules whose microsymbionts appear to be largely limited to
*Bradyrhizobium* (Fonseca et al., 2012; Parker, 2015; Ardley &
Sprent, 2021) and which are mostly confined to the tropics and
subtropics where bradyrhizobia are dominant and widespread
(Parker, 2015; Meng et al., 2019).

Despite the significantly lower rates of loss of nodulation in
mimosoids with SYM-type nodules compared to FTs in the
remaining Caesalpinioideae, SYM-type nodulation is not imper-
vious to evolutionary loss. Our reconstruction documents five
losses of nodulation within the Mimosoid clade: on the branches
subtending the *Amblyogonocarpus* + *Adenanthera* clade, and the
genera *Parkia*, *Cylicodiscus*, *Newtonia* and *Zapoteca* (Fig. 1;
Table S3). These losses in ancestrally SYM-type lineages suggest
that even the advantages conferred by stricter SYM-type compart-
mentalization can be insufficient to prevent losses of nodulation.
Within the genus *Senegalia*, a small subset of species have appar-
ently lost the ability to nodulate (Sprent, 2001, 2009). Current
phylogenies imply that nodulation has been lost in species of each
of the two main subclades within *Senegalia*, offering a useful sys-
tem for understanding environmental or other factors associated
with loss of nodulation.

Conclusions

The evolution of the symbiosome in species-rich nodulating legume
lineages offers a compelling explanation for the well-known but
poorly understood highly uneven distribution of nodulating species
richness across the N₂-fixing clade. While nodulation has been sug-
gested as a possible key innovation underpinning the evolutionary
success of legumes, our results suggest that it was adoption of SYM-
type nodules and the innovation of the symbiosome that under-
pinned the stabilization of N₂-fixation and potentially contributed
to massive diversification of species within Caesalpinioideae and
Papilionoideae, the two most diverse and geographically widespread
subfamilies of legumes. Furthermore, the greater propensity of the
FT-type nodule to be secondarily lost and for SYM-type lineages to
persist and diversify provides a potent example of the long-term
evolutionary benefits and outcomes of stricter compartmentalization
in symbiotic cooperation, as suggested by Chomicki *et al.* (2020).

We show that the grade of Caesalpinioideae lineages subtend-
ing the Mimosoid clade is a hotspot of evolutionary transitions
between phylogenetically intermingled nodulating and nonnodu-
lating lineages (Fig. 1), including two independent transitions
from FT- to SYM-type nodules as well as numerous losses of
nodulation. The phylogeny and detailed evolutionary trajectories
of nodulation and nodule anatomies presented here provide a
robust framework for comparative genomic analyses of FT and
SYM nodulating and nonnodulating lineages across Caesalpin-
ioidiae. This includes *Chamaecrista*, the most species-rich genus
in the nonmimosoid grade of Caesalpinioideae, and the only
nodulating genus in the subfamily which contains species with
both SYM-and FT-type nodules (Naisbitt *et al*., 1992), thus rep-
resenting a second possible transition from FT to SYM.
## Acknowledgements

We thank Gwilym Lewis for help with identification of plants, the Millennium Seedbank for supplying seeds and colleagues who have helped sample material used in this study. Jordan Price is thanked for his expert advice on actinorhizal symbioses. This work was supported by the research project Engineering Nitrogen Symbiosis for Africa (ENSA), which is funded by a grant to the University of Cambridge by the Bill & Melinda Gates Foundation and the Foreign, Commonwealth & Development Office (FCDO), the Swiss National Science Foundation (grants 310003A_156140 and 31003A_182453/1 to CEH), CNPq grant 312125/2020-8 to SMF, and Fundação de Amparo à Pesquisa do Estado da Bahia (FAPESB, grant APP0037/2016 to DC).

## Author contributions

EJMK, JJR, CEH and EKJ designed the study and interpreted the results, JJR carried out the phylogenetic analysis, SMF, EKJ, EG, KMW and YP performed anatomical analyses, SMF, EKJ, EG, DC, GKDA, J Akomatey, NT, HSG, YP, MM, NT, PS, HCL and CEZ sampled nodules in the field, and SMF, EJMK, JJR, EKJ, J Ardley, DC, EG, YP, JIS and CEH wrote the paper.

## ORCID

Julie Ardley ID https://orcid.org/0000-0001-9938-606X
Domingos Cardoso ID https://orcid.org/0000-0001-7072-2656
Sergio M. Fariade ID https://orcid.org/0000-0001-5066-046X
Hukam S. Gehlot ID https://orcid.org/0000-0001-7447-4960
Eduardo Gross ID https://orcid.org/0000-0002-1057-0432
Colin E. Hughes ID https://orcid.org/0000-0002-9701-0699
Euan K. James ID https://orcid.org/0000-0001-7969-6570
Erik J. M. Koenen ID https://orcid.org/0000-0002-4825-4339
Haroldo C. Limade ID https://orcid.org/0000-0003-2154-670X
Marta Maluk ID https://orcid.org/0000-0002-5038-0094
Yves Prin ID https://orcid.org/0000-0002-3706-0045
Jens J. Ringelberg ID https://orcid.org/0000-0003-0567-5210
Nisha Tak ID https://orcid.org/0000-0002-3097-4119
Neung Teaumroong ID https://orcid.org/0000-0002-2788-9458
Kathryn M. Wright ID https://orcid.org/0000-0002-5815-0808
Charles E. Zartman ID https://orcid.org/0000-0001-8481-9782

## Data availability

The data that support the findings of this study are available in the Supporting Information of this article.

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**Supporting Information**

Additional Supporting Information may be found online in the Supporting Information section at the end of the article.

**Fig. S1** Symbiosomes are standard in nodules of caesalpinioids from the Mimosoid clade.

**Fig. S2** Fixation threads (FTs) are standard in nonmimosoid grade Caesalpinioid nodules.

**Fig. S3** Evolutionary trajectory of nodulation and nodule type when transitions between all nodulation states are allowed. Methods and legend otherwise as for Fig. 1.

**Fig. S4** Evolutionary trajectory of nodulation and nodule type when taxa with missing data have been assigned equal weight to nodulation and nonnodulation states.

**Fig. S5** Evolutionary trajectory of nodulation and nodule type when two additional losses of nodulation within *Senegalia* are included.

**Notes S1** References for Table S2.

**Table S1** Caesalpinioideae and outgroup taxa used in the time-calibrated phylogeny depicting evolutionary trajectories of nodulation and nodule type.

**Table S2** Occurrence of fixation threads (FTs) and/or symbiosomes in nodules from Caesalpinioideae (Mim, belongs to the Mimosoid clade) extracted from the literature and from the unpublished observations of the authors.

**Table S3** Type, location and age of transitions in nodulation status as depicted in Fig. 1.

**Table S4** Summary of rates of losses of nodulation per million years across analyses.

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