www.sciencemag.org/cgi/content/full/326/5956/1120/DC1

Supporting Online Material for

Symbiotic Nitrogen Fixation in the Fungus Gardens of Leaf-Cutter Ants

Adrián A. Pinto-Tomás, Mark A. Anderson, Garret Suen, David M. Stevenson, Fiona S. T. Chu, W. Wallace Cleland, Paul J. Weimer, Cameron R. Currie*

*To whom correspondence should be addressed. E-mail: currie@bact.wisc.edu

Published 20 November 2009, Science 326, 1120 (2009)

DOI: 10.1126/science.1173036

This PDF file includes:

Materials and Methods

SOM Text

Figs. S1 to S10

Tables S1 to S6

References and Notes

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Materials and Methods Study organisms. Leaf-cutter ant colonies were collected in Argentina (Misiones, 2003), Panama (Canal Zone, 2003 and 2006), and Costa Rica (La Selva [Heredia] and Palo Verde [Guanacaste] Biological Stations, 2006; Corcovado National Park [Puntarenas], 2008). For colonies collected in Costa Rica, microbial isolations were conducted at the laboratories of each research station. Colonies were transported to the University of Wisconsin-Madison, where they were maintained inside plastic polystyrene boxes connected by plastic tubes and placed individually over mineral oil islands to help prevent the potential horizontal transfer of microbes by mites. Ants were fed local foliage (Northern Pin Oak, Quercus ellipsoidalis, and Norwegian Maple, Acer platanoides) three times per week and provided with water for moisture once a week. N content measurement. Five core samples were taken from each of five Atta cephalotes colonies by slowly introducing an inverted sterile 15 mL conical centrifuge tube from top to bottom of the fungus garden. Three of these samples were carefully sorted into fungus garden, ant workers and ant brood. Duplicate refuse dump samples were similarly obtained. Leaves from six Madison, WI trees (three A. platanoides and three Q. ellipsoidalis) regularly used to feed these colonies were also collected. Samples were stored overnight at -20ºC, dried at 55ºC for 48h and weighed. N content was determined by nitrogen combustion (Dumas) analysis, employing either a varioMAX CN (Elementar Analysensysteme, Hanau, Germany) for small volume samples (ant workers and brood) or a LECO FP-2000 (Leco, St. Joseph, MI) for all other samples. One-way ANOVA statistical tests were performed with SAS software, employing the GLM procedure (SAS Institute Inc., Cary, NC). Acetylene reduction (AR) assay. AR with leaf-cutter ant colonies was conducted by placing triplicate one gram (wet weight) samples of each fungus garden, ant workers and ant brood in stoppered 9-ml vials containing 90% air and 10% acetylene (v/v). We also conducted AR with samples from leaves used to feed the ants and with samples of refused material. Samples were incubated at 28ºC for 48 h, and ethylene production was measured periodically by gas chromatography (S1). To standardize AR measurements by dry weight, the samples were dried at 65ºC for 48 h and weighed. Statistical tests were performed with SAS software, employing the GLM procedure (SAS Institute Inc). AR was conducted on samples from 35 leaf-cutter ant colonies from eight different species collected across three different ecosystems, as summarized in Table S1. For AR conducted in La Selva, Costa Rica, each test was performed as described above and gas samples were stored in sterile Vacutainer tubes (Becton-Dickinson, Franklin Lakes, NJ) until transport to Madison, where ethylene production was measured by gas chromatography. Isolation of N2-fixing bacteria. All incubations were performed at 28ºC. Five pieces (~1 mm3) of fungus garden from each leaf-cutter ant colony (n = 80) were placed in duplicate tubes containing 10 mL of N-free media (S2), and incubated in agitation for 4 h. Then 100 µL of 1:10 dilutions were inoculated in duplicate N-free media plates (2% noble agar) and test tubes containing semi-solid N-free media (0.5% noble agar) and incubated for

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~120h. Growth pellicles in semi-solid media were streaked to N-free plates. Individual colonies in N-free media were transferred to nutrient agar. For AR assays, pure cultures were inoculated in nutrient broth and incubated in agitation for 24 h. Bacterial pellets were washed 3 times and re-suspended in 1 mL washing buffer (S3). 100 µL of this suspension were inoculated in 25 mL test tubes containing 15 mL of semi-solid N-free media. Tubes were incubated for 72 h, stoppered with rubber plugs and injected with 1 mL of acetylene. Ethylene production was measured periodically by gas chromatography. Azospirillum sp. was used as positive control. DNA extraction, PCR amplification and sequencing. Fungus garden, ant tissue, fungal cultivar and Pseudonocardia samples were subjected to CTAB-based DNA extraction protocols (S4, S5). DNA was cleaned with DNA Cleanup & Concentrator (Zymo Research, Orange, CA). PCR reactions were performed with GoTaq® Green Mastermix (Promega, Madison, WI) in MJ Research PTC-200 thermocyclers. To amplify nifH we used universal primers (S6): nif-Fo (5′-AAAGGYGGWATCGGYAARTCCACCAC-3′) and nif-Re (5′-TTGTTSGCSGCRTACATSGCCATCAT-3′) using the stringent PCR conditions described by Widmer and collaborators in order to reduce the emergence of falsely positive amplifications (S7), including the following cycling conditions: 95ºC/5min, 40 cycles of 94ºC/11s, 92ºC/15s, 54ºC/8s, 56ºC/30s, 74ºC/10s and 72ºC/10s, and final extension for 10 min/72ºC. PCR products were cleaned with ExoSAP (USB Corporation, Cleveland, OH) and cloned with the TOPO TA Cloning Kit (Invitrogen, Carlsbad, CA). Clones showing inserts of expected size (~450bp) were randomly selected for sequencing. For N2 fixers in pure culture, we performed colony-PCR. The 16S rDNA gene was amplified using universal Domain Bacteria primers (S8) with the following program: 95ºC/4min, 30 cycles of 94ºC/45s, 51ºC/50s and 72ºC/140s, and a final extension period of 5min/72ºC. We performed multilocus sequence typing of housekeeping genes gapA, icdA and mtlD for confirmed N2 fixers using the primers and conditions described by Ma and colleagues (S9). PCR products were cleaned with ExoSAP prior to preparing sequencing reactions with the BigDye reaction mix (Perkin-Elmer Corp., Foster City, CA). Excess dye terminators were removed using CleanSEQ™ reaction (Agencourt Bioscience, Beverly, MA) and loaded into an Applied Biosystems 3700 automated DNA sequencing instrument. Sequences were deposited in GenBank under accession numbers FJ593730-FJ593840. Phylogenetic analyses. Nucleotide sequences were analyzed and manually edited using Sequencher 4.5 (Gene Codes Corporation). 16S rDNA sequences were aligned using the NAST algorithm in arb (S10) with an rDNA database containing the Greengenes 16S library (S11) (accessed: 01/15/2009). The alignment of multilocus sequences was performed by concatenating each sample set of sequences and then aligning them using MUSCLE (S12). Phylogenetic analyses for both 16S rDNA and multilocus sequences were conducted with MrBayes v. 3.0b4 (S13). Each analysis consisted of four independent chains, one cold and three incrementally heated (T=0.2), starting from random trees, and was run for five million generations with trees sampled every 100 generations to calculate posterior probabilities for each branch. The majority rule consensus tree was calculated after removing the first 500 trees corresponding to the burn-in period estimated according to the log-likelihood curve. The General Time

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Reversible (GTR) model was used with priors for the substitution rates set to a flat distribution. 15N-enrichment experiments. Duplicate (paired) sub-colonies weighing 10 g were placed in individual airtight containers (Fig. 3A) consisting of two chambers connected by plastic tubing and an airtight valve. The fungus garden chamber included a CO2 trap (modified cell culture vial with vented cap containing 4 g of soda lime) and a moisture device (pierced plastic cylinder containing moist cotton). The feeding chamber included a PVC block to reduce volume and provide space for the ants to dump their waste (between the block and the container wall). Each container was subjected to a brief vacuum and its atmosphere was replenished with a mixture of 80% 15N2 (Cambridge Isotope Laboratories, Andover, MA) and 20 % O2 (treatment group) or regular air (~80% 14N2, control group), using 500 mL airtight syringes (Hamilton Co, Reno, NV). Paired sub-colonies were fed three equivalent fragments of the same leaves (A. platanoides). Feeding and atmosphere exchange were performed twice every week. Samples were frozen overnight, carefully sorted into fungus garden, ant workers, and ant brood, dried for 48 h at 60ºC and homogenized. Ant workers’ gasters were cut off at the petiole prior to homogenization, since undigested food can interfere with isotope measurements (S14). To analyze N isotopic ratios, sub-samples of ant workers or brood (~8 mg), and fungus garden (~12 mg) were placed in quartz tubes (7mm i.d. x 9mm o.d. x 25 cm), which were charged with ≈5 g of CuO, ≈ 0.5 g of Cu and ≈ 250 mg of Ag, and then evacuated, flame sealed, and combusted at 800ºC for 6 hours. Following this, the tubes were placed on a high vacuum line and distilled through two -78°C traps and one -196°C trap and the N2 was then trapped on 5 Å molecular sieves at -196°C. The sieves were heated to 250°C to drive off the N2 gas which was then analyzed using a Finnegan MAT Delta E Isotopic Ratio Mass Spectrometer (IRMS). The IRMS determines the isotopic ratio (percentage of 14N and 15N in each sample) compared to a known standard to give a δ value:

δ = 1000[{(15Nsample/14Nsample)/(15Nstandard/14Nstandard)}-1] The statistical significance of 15N enrichment was evaluated through paired t-tests comparing the δ values of control and experimental sub-colonies. Additionally, we determined 15N natural abundance in leaves of five A. platanoides and five Q. ellipsoidalis trees local to Madison, WI that we regularly employed to feed the leaf-cutter ant colonies. Leaf samples were stored at -20ºC in airtight bags for two months and then measured as this is the routine procedure used to feed the leaf-cutter ant colonies during winter. Sequencing and identification of the nif cluster from a Klebsiella and Pantoea isolate. The nif cluster from isolates Klebsiella variicola At-22 and Pantoea sp. At-9b were sequenced at the U.S. Department of Energy Joint Genome Institute (S15). Sequencing of the nif clusters was performed using 454 pyrosequencing (S16) (454 Life Sciences). General aspects of library construction, sequence assembly, and quality assessment can be found at http://www.jgi.doe.gov. Automated annotation of the nif clusters were identified and performed at the U.S. Department of Energy Oak Ridge National Laboratory, and details regarding this automated annotation process can be found at

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http://genome.ornl.gov. To identify the nif pathway in both isolates, the nif cluster from the complete sequenced genome of Klebsiella pneumoniae 342 (loci: KPK_1696 – KPK 1714, GenBank Accession: CP000964.1) was used as a scaffold for classifying each isolates’ respective cluster, and confirmation of these pathways was performed using BLAST (S17). The full nucleotide and predicted protein sequences for the nif clusters from both Klebsiella variicola At-22 and Pantoea sp. At-9b have been deposited into GenBank under the Accession numbers GQ342603 and GQ342604, respectively. Real-Time PCR. A total of 6 g of fungus garden material was obtained from each of 8 lab-maintained Atta cephalotes colonies. Each sample was separately suspended in 40 mL of cold extraction buffer as previously described (S18) with the following pre-processing steps. Each sample was vortexed to dislodge bacteria, and filtered by squeezing through 4 layers of cheesecloth. The filtrate was then centrifuged at 8000 g for 20 min, the supernatant poured off, and the cells washed with another 10 ml cold extraction buffer. Finally, cells were re-suspended in about 800 µl of buffer. For controls, DNA was extracted as described previously (S18) from pure cultures of two isolates obtained from leaf-cutter ant fungus gardens, Klebsiella variicola At-22 and Pantoea sp. At-9b. Resultant DNA was treated with DNase-free RNase, ethanol-precipitated, and finally resuspended in TE. Total DNA was quantified spectrophotometrically, and diluted to a 10 ng/µl working concentration. Real-time PCR was performed as previously described (S18). All samples were run in quadruplicate, with their respective primers, and the efficiency was confirmed using pure culture isolates for Klebsiella and Pantoea. Because no nifH-containing isolate in the genus Burkholderia was obtainable from leaf-cutter ant fungus gardens, Klebsiella standards were used to evaluate Burkholderia, under the assumption that the efficiency was the same for both. In addition, the same fluorescence crossing threshold (Ct) value was selected across all PCR experiments. Samples were then run with the universal nifH primers (total nifH), Klebsiella-specific nifH primers, Pantoea-specific nifH primers, and Burkholderia-specific nifH primers. Both positive and negative (no template) controls were also run. Primer sets for each test were as follows: Universal nifH: nif-Forward-qPCR: 5’-GGWATCGGYAARTCCACCAC-3’

nif-Reverse-qPCR: 5’-CSGCRTACATSGCCATCAT-3’ Klebsiella-specific nifH: KleUniNifH1F: 5’AAGAAAGTGATGATCGTCGGCT-3’ KleUniNifH1R: 5’-CCTCGACCGAGCCCACTT-3’ Pantoea-specific nifH: PanUniNifH1F: 5’-AAGAAGGAGCCTACGTACCCG-3’, PanUniNifH2F: 5’-AGAAGGTGCCTACGTGCCC-3’, PanUniNifH3F: 5’-AAGAAGGTGCCTACGTACCCG-3’, PanUniNifH4F:5’-AAGAAGGTGCCTACGTACCTGATC-3’ PanUniNifH1R: 5’-CGGATGGGCATGGCAA-3’ PanUniNifH2R: 5’-TCACGGATAGGCATGGCAA-3’ Burkholderia-specific nifH: BurUniNifH1F: 5’-TCGTACGACGTGCTCGGC-3’ BurUniNifH1R: 5’-TCCATCGTGATCGGCGTC-3’ Transmission Electron Microscopy (TEM) Eight samples of Atta cephalotes fungus gardens were fixed in Karnovsky solution (2.5% glutaraldehyde, 2.0% paraformaldehyde) in 0.1 M phosphate buffer, pH 7.4. Samples were included in 2.0% agarose (Sigma) in

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phosphate buffer and cut in 1.0 mm3 sections. The samples were post-fixed with 1% osmium tetroxide. The preparations were dehydrated in ethanol, then propylene oxide was used as intermediate solvent, before finally embedding the samples in Spurr resin. Ultra-thin sections (60-70 nm) were obtained with a Reichert Ultracut’s ultramicrotome (Leica Wien, Austria), stained with 4% uranyl acetate and 2% Sato’s triple lead and observed with Hitachi H-7.100 or H-7.000 transmission electron microscopes working at 100 kV. Supporting Text 1. Nitrogen limitation and enrichment in leaf-cutter ant colonies Leaf-cutter ant workers use fresh plant substrate to cultivate their mutualistic fungus. Although the substrate collected includes fruits, seeds, flowers and stipules, the vast majority is fresh leaves (S19). Thus, the ants, through the association with their fungus garden acting as an ancillary digestive system, are functionally herbivores. As a food source, leaves have a much lower ratio of nitrogen to carbon (N:C) than is required by insects. The N:C ratio in leaves of woody plants ranges between 1:100 to, at best, 1:20 (S20), whereas the N:C ratio is around 1:10 in insect tissues and 1:6 in chitin, the main component of the exoskeleton (S21). This leads to the prediction that leaf-cutter ant colonies are growth limited by N. There are a number of ways the leaf-cutters could supplement N, including acquisition through mineralization from the soil by the fungus garden, the ants feeding on additional N-rich food sources, and/or colonies receiving indirect access to atmospheric N2 through symbiotic associations with N2-fixing bacteria. Leaf-cutter ant fungus gardens are vertically stratified. The newest section at the top is where workers integrate new leaf substrate, while the oldest part is at the bottom of the garden. As part of the regular turnover of garden material, the ants remove partially degraded plant tissue (i.e., exhausted fungus garden) from the bottom section and place it in refuse dumps located in subterranean chambers or in conspicuous external mounds (e.g., Atta colombica). This facilitates research on the nitrogen balance of leaf-cutter ant colonies by allowing a comparison between N input, in the form of leaves, and N output, in the form of spent garden substrate removed by workers. If N is a growth limiting resource for leaf-cutter ant colonies, then it should be preferentially depleted from leaf substrate resulting in discarded substrate being enriched for C relative to N. In contrast to this prediction, studies by Haines (S22) showed that N is enriched in the refuse dump when compared with the harvested leaf material. A net N gain in the refuse material compared to harvested plant tissue was also reported for Acromyrmex lundi (S23). Similarly, a long-term study with A. colombica in Panama revealed that N concentration is 26 times higher in the colony refuse than in the surrounding leaf-litter (S19). The “nutritional enrichment” of the material discarded by the leaf-cutters partly explains why their dumps play an important role in nutrient cycling within tropical ecosystems, as illustrated by the significant enhancement of fine root production within the dump when compared to the general forest floor (S19, S22, S24). In addition, greater plant diversity is

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observed within abandoned A. cephalotes nests (with underground refuse dumps) than in the adjacent forest (S25). Taken together, these results suggest that the ants obtain additional N from sources other than leaves, potentially through associations with N2 fixers. However, these studies were all conducted in the field, where the observed N enrichment could be due to mineral uptake from the soil, or by workers foraging on high N food sources to compensate for the limited N content of leaf substrate. Therefore, we analyzed the N content of laboratory maintained A. cephalotes colonies, where N input supplementation is limited to just N2 fixation. These lab colonies were only fed leaves from maple (Acer platanoides) and oak (Quercus ellipsoidalis) trees local to Madison, WI. This feeding occurred three times per week, and not “ad libitum”, so compensatory feeding and exploitation of additional N rich substrates were both prevented. The N content of both types of leaves in July 2007, when this experiment was conducted, was 2.44% ± 0.19% s.e.m and 2.43% ± 0.10 % s.e.m, for maple and oak, respectively. Our N-content analysis clearly shows that, under controlled laboratory conditions, N is enriched as materials flow through leaf-cutter ant colonies (Fig 1C, main text). As expected, worker ants and ant brood have higher N content, because their tissue and exoskeleton both contain a higher N:C ratio, corresponding with selective N retention during metabolism. Both live and dead ants were removed from the fungus garden and refuse material before measuring N content, therefore enrichment in the refuse dump was not due to the presence of insect tissue. Our results confirm previous field studies, and, by preventing acquisition of additional N through foraging or mineralization, they provide indirect evidence for the occurrence of N2 fixation within leaf-cutter ant colonies. It is possible that the consistent N:C enrichment observed in lab and field colonies of leaf-cutter ants is a result of excess C utilization by the ants and their cultivated fungus (i.e., respiration by the ants and fungi results in excess use of C relative to N). Although excessive respiration of C is likely a partial explanation for the N:C ratios observed in these studies, it is unlikely to account for the full extent of N enrichment observed. This conclusion is based on the fact that leaf-cutter ant colonies experience intense interspecific and intraspecific competition for resources in Neotropical forests, which has selected for the energetic optimization of tasks related to plant material harvesting and fungus garden cultivation and management (S26, S27). This selection pressure should also extend to efficient use of carbon in leaf-cutter ant colonies. Nevertheless, despite laboratory evidence for N enrichment through N2 fixation, definitive evidence for this phenomenon occurring in leaf-cutter ant colonies requires a combination of functional tests for N2 fixation and the demonstration that fixed N is incorporated into fungus garden and ant biomass. 2. Demonstration of N2-fixing activity within leaf-cutter ant colonies by acetylene

reduction assays N2 fixation in biological systems is sometimes tested by attempting to detect the presence of N2-fixing bacteria and/or the genes responsible for this process. This however is insufficient for several reasons. First, bacteria belonging to genera known to fix N2 are

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ubiquitous in nature (S28, S29). Second, nitrogenase genes, such as nifH, are so widespread in environmental sources that they are known to contaminate PCR reagents (S28). Third, the process of N2 fixation is energetically costly, and therefore highly regulated at the transcriptional level by a complex regulatory network integrating several environmental cues, thus the nitrogenase enzymatic complex may only be active at specific times or niches (S30). Instead, it is recognized that conclusive evidence for N2 fixation must include positive functional tests (S31, S32). Acetylene reduction (AR) is an excellent assay to evaluate the occurrence of N2 fixation, because positive results imply the presence of a functional nitrogenase enzyme reducing acetylene’s triple bond (analogous to dinitrogen triple bond) to produce ethylene (S1). To address our hypothesis that leaf-cutter ants have symbiotic associations with N2 fixers to supplement their N budget, we conducted extensive AR assays. We detected AR in all leaf-cutter ant fungus garden tested (n = 35). In addition, positive results were obtained for at least two ant colonies from five species of Acromyrmex sp. and three species of Atta sp. (Table S1). The detection of N2 fixation within the fungus gardens of leaf-cutter ants by AR included: i) A. cephalotes colonies tested directly in the field in Costa Rica (Fig. S2.B); ii) colonies of leaf-cutter ants maintained in the lab for less than 1 month after field collection (see Table S1, colonies collected in 2008); and iii) Leaf-cutter ant colonies maintained in the lab for more than 3 years (Fig. S1). AR tests on colonies from this latter group showed that activity levels remained relatively constant over a 2 year period (Fig. S1). This finding indicates that N2 fixers maintain a continuous presence within the ant’s fungus garden, because the leaves used to feed the ants in the laboratory do not exhibit positive AR, and apparently do not harbor N2-fixing bacteria (see below). No significant AR activity was detected in worker ants, ant brood, or in leaves used to feed the leaf-cutter ant colonies in the laboratory (Fig. 1D, Fig. S3). Having revealed the occurrence of nitrogenase activity within leaf-cutter ant colonies, we performed additional assays to further localize this activity within the fungus garden of eight colonies of Atta sp., both in the laboratory and directly in the field. We conducted AR from three different sections of the fungus garden, including: i) the top, which is the youngest section where fresh leaf material is integrated; ii) the middle region where the ant brood and most of the fungus-produced feeding structures (gongylidia) are located; and iii) the bottom and oldest section, representing partially exhausted plant and fungus tissue. We found AR activity to be highest in the middle section of the fungus garden (Fig. S2). Because this region is where most feeding structures are located (S33), this suggests that N2 fixation directly benefits the ants. Further, it provides additional evidence that N2 fixation is not a result of N2-fixing bacteria inoculated in the fungus garden with the leaf substrate, since the newest leaf material is located at the top of the fungus garden. Because the nitrogenase enzyme is sensitive to oxygen, and our AR assays were conducted without removing O2 from the assay vials, we compared the activity of different fungus garden samples under aerobic and anaerobic conditions. Our results show that fungus garden samples exhibit similar AR activity in both cases (Fig. S4), indicating that O2 removal is not necessary to obtain positive N2 fixation in leaf-cutter ant

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fungus gardens. Further, this indicates the existence of specific sites within the fungus garden matrix that allow N2 fixation in the presence of O2. Taken together, our AR results demonstrate the occurrence of biological N2 fixation within the fungus garden of leaf-cutter ant colonies. This conclusion is supported by our findings that: i) all fungus gardens sampled from eight different species of leaf-cutter ants exhibited positive AR; ii) AR levels were consistent over a two year period in laboratory maintained colonies; iii) positive AR activity was detected directly in field colonies; and iv) AR activity was highest in the middle of the fungus garden, where ant feeding is concentrated. 3. 15N2-enrichment experiments establish that fixed N is incorporated into leaf-

cutter ant biomass If leaf-cutter ants obtain a direct benefit from N2-fixing bacterial symbionts, then fixed N should be incorporated into the ants’ biomass. We tested for the integration of fixed N in the ants by employing stable isotope enrichment experiments. Nitrogen has two natural isotopes, 14N (~99.6% of the N2 in the atmosphere) and the less abundant 15N. The proportion of both isotopes can be measured in the tissue of a given organism and the ratio of 15N to 14N is termed δ 15N [per mil (‰)]. In our experiments, we compare the δ 15N values for paired sub-colonies of leaf-cutter ants, one maintained in a normal atmosphere and the other maintained in a 15N2-enriched atmosphere where we replaced 14N2 with

15N2. Then by monitoring for an increase in 15N in both the fungus garden and the ants in the enrichment treatment relative to the control, we track the potential passage of fixed N in the system (i.e., any observed increase in δ 15N reflects input from N2 fixation). The basis of our experiments is that, if N2 fixers contribute a significant amount of N to leaf-cutter ant biomass, then ants from sub-colonies maintained in an atmosphere composed mostly of 15N2 will have a higher δ 15N value than those grown in normal atmospheric conditions. To conduct this experiment, we designed a novel airtight apparatus to allow for the exchange of a normal atmosphere for one enriched in 15N2 (Fig. 2A). The addition of a CO2 trap and a moisture device allowed the ants to normally function for ~72 hours without any atmosphere exchange. Several preliminary trials were run to optimize the experimental conditions and to determine the length of time these sub-colonies were healthy and stable. We found that, despite optimizing the experimental conditions, the health of the majority of the colonies deteriorated after two weeks. As a result, we limited the length of our main experiment (see text) to a two week period. Our experiments reveal that fixed N is incorporated into the fungus garden and the ants (see Fig. 2B-C, Tables S3 and S4). The level of 15N enrichment (δ 15N increase in treated versus control sub-colonies) was consistently higher in the fungus garden (week 1 = 0.90 ‰, week 2 = 1.50 ‰) than in the worker ants (week 1 = 0.47 ‰, week 2 = 0.64 ‰) and the ant brood (week 1 = 0.30 ‰, week 2 = 0.26 ‰) (Fig. S5, evaluated by one-way ANOVA, week 1: F2,21 = 7.66, P = 0.0032; week 2: F2,18 = 6.68, P = 0.0068), thus confirming that the fungus garden is the main site for N2 fixation within leaf-cutter ant colonies. In addition, some of our colonies were still healthy and performing normal tasks one month after the

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start of pilot experiments, and significant 15N enrichment was also found in them (Fig. S6). 4. Isolation and identification of symbiotic N2-fixing bacteria associated with leaf-

cutter ants To isolate and identify the symbiotic N2-fixing bacteria associated with leaf-cutter ants we focused on the fungus garden of 80 colonies from eight species of leaf-cutters. Specifically, we performed isolations using a traditional enrichment technique with N-free media under aerobic conditions (S2). Azospirillum sp., a well-known free-living diazotroph, was used as a positive control for all tests. Colony forming units obtained in N-free media were isolated in pure culture, identified through 16S rDNA sequencing, and evaluated for N2-fixing capability by AR and PCR amplification of the nifH gene. Only isolates in the genera Klebsiella and Pantoea were confirmed to fix N2 (Fig. 3A, Table S5). There are several potential explanations for the fact that isolates obtained in N-free media belonging to the other 15 genera did not show positive AR, including i) growth on N-free media being a result of interactions with true N2 fixers; ii) these isolates may require complete removal of oxygen from the AR test tubes in order to fix N2; and iii) isolates may posses an unusual nitrogenase complex that does not reduce acetylene to ethylene, as has been shown for some actinobacteria (S34). Independent of these considerations, N2 fixers in the genera Klebsiella were the most frequently isolated from fungus gardens sampled directly in the field or immediately after field collection (69% of evaluated colonies where shown to be colonized by this symbiont, n = 58). Furthermore, we were able to consistently isolate Klebsiella from leaf-cutter ant colonies maintained in our laboratory in Madison, WI several months after their field collection. Klebsiella was not isolated from the leaves used to feed the ants in Madison, which indicates that these symbionts are not contaminants from the plant material added to the colony, but instead form a consistent association within the fungus gardens of leaf-cutter ants. Given their high N2 fixation rates, the stability of their association with leaf-cutters, the fact that they form a specialized monophyletic clade in our phylogenetic analyses (see below), and their ability to establish symbiotic interactions with other types of insects (S31, S35, S36), isolates in the genus Klebsiella are likely one of the main N2-fixing symbionts colonizing the fungus gardens of leaf-cutter ants. The other group of N2 fixers we found associated with leaf-cutter ants belongs in the genus Pantoea. These bacteria also fix N2, although at a slower rate than the Klebsiella symbionts (Fig. 3A, Table S5). Pantoea isolates have also been identified as symbionts of different insect species, including locusts, thrips and phylloxera (S37-S39). Interestingly, there appears to be an interaction between Pantoea and Klebsiella symbionts associated with the locust Schistocerca gregaria, whose gut colonization by Pantoea is enhanced by the presence of Klebsiella symbionts (S40). Furthermore, the nifH gene from both groups of leaf-cutter ant-associated N2 fixers is closely related to the nifH gene of Klebsiella sp. (Fig. S7), highlighting the potential for horizontal gene transfer between these two genera.

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5. Phylogenetic analyses of N2-fixing symbionts associated with leaf-cutter ants. We initially identified all fungus garden microbes isolated in N-free media by sequencing their 16S rDNA gene, and then constructed a phylogeny for the two main groups of confirmed N2 fixers associated with leaf-cutter ants (Fig. S8). The 16S rDNA fragment (1400bp) was identical for all confirmed N2 fixers in the genus Klebsiella, and was a complete match to the type strain of Klebsiella variicola. In our 16S phylogeny, the closest relative for the other group of leaf-cutter ant-associated N2 fixers was Erwinia cypripedii (currently Pectobacterium cypripedii). However, phylogenies for bacteria based exclusively on 16S genes lack resolution below the genus level, and even the family level in some groups (S41); hence, they are not sufficient to resolve specific associations between bacteria and their multicellular hosts. Therefore, to further explore the taxonomy and specificity of the leaf-cutter ant–associated N2 fixers we performed multilocus sequencing of three housekeeping genes (gapA, icdA, and mtlD). These genes have been successfully employed to construct robust phylogenies that correlate well with phenotypic characterization for bacteria in the Family Enterobacteriaceae (S9, S42). We were able to obtain and sequence the type strain for Klebsiella variicola, but otherwise our phylogenetic analyses were limited by the available sequences of the selected housekeeping genes in public genome databases. We took advantage of a recent phylogeny of Erwinia and their close relatives reported by Ma and collaborators (S9). This is significant, as our 16S rDNA phylogeny indicates that the second most common N2 fixer group found associated with leaf-cutter ant colonies belong to the genus Erwinia, which has undergone an extensive phylogenetic revision in the last few years, including some species being placed into at least 4 different genera (S9). Indeed, our multilocus phylogeny indicates that this group of ant–associated fixers does not belong in any of the clades of plant pathogens, including the genera Erwinia and Pectobacterium, but rather belong to the genus Pantoea (Fig. 3B). Our phylogenetic analyses of ant-associated N2 fixers in the genus Klebsiella reveal that they are closely related to the plant-associated species Klebsiella variicola (S43). It is possible that the origin of Klebsiella in the ant system is through continuous re-acquisition of these bacteria from the environment, since plant tissue turnaround in leaf-cutter ant colonies is constant and continuous. Although we currently can not rule out this possibility, it appears unlikely. This is based on the bacteria maintaining a continuous presence in the ants’ fungus garden over time in lab colonies; which lack an apparent source of inoculum of K. variicola in the leaves provided (see above); the presence of these bacteria in leaf-cutters from broad geographic areas; and our finding that Klebsiella isolates are associated with diverse species of leaf-cutter ants, including those from different ecosystems. Instead, it is likely that the grouping of K. variicola within the leaf-cutter ant–associated Klebsiella clade is based on a lack of resolution due to the slow-evolving nature of these housekeeping genes. Future studies will likely require a phylogenomic approach, comparing the genomes of these bacteria. Nevertheless, our AR and 15N-enrichment results indicate that in the unlikely event that the source of the N2 fixer is through leaf material in tropical ecosystems, it appears that the ants have

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evolved to utilize these otherwise free-living bacteria, maintaining them in their garden to supplement their N budget. 6. Estimating the amount of symbiotic N2 fixation in leaf-cutter ant colonies and its

potential ecosystem impact In the Neotropics, leaf-cutter ants are arguably a keystone species, helping shape the ecosystems in which they occur (S44). Through their prolific pruning of vegetation they help structure surrounding plant communities, stimulating new plant growth, breaking down plant material, and enriching the soil (S45). Perhaps equally important, the leaf-cutters play a major role in nutrient cycling within these ecosystems (S46, S47). Previous studies have established that leaf-cutter ant colonies concentrate N. For example, Wirth and collaborators (S19) reported that N concentration is 26 times higher in A. colombica refuse dumps than in the surrounding leaf-litter. Our findings that symbiotic N2 fixation occurs in the fungus gardens of leaf-cutter colonies, and that fixed N is integrated into ant biomass, indicate that the observed soil nutritional enrichment is derived, at least partially, through biological N2 fixation. In this section we provide estimates of the impact of this phenomenon, both at the colony and at the ecosystem level.

To estimate the impact of symbiotic N2 fixation on leaf-cutter ant colonies we first employ a method based on the natural abundance of the heavier N isotope, 15N (measured as δ 15N [per mil (‰)], see section 3 above). Specifically, the contribution of symbiotic microorganisms towards the N budget of their hosts can be estimated by comparing the δ 15N of the host with the δ 15N of their diet. In catabolic reactions the lighter isotope (14N) is lost at a greater rate, resulting in progressive enrichment of 15N in each trophic level transfer. Furthermore, as mentioned previously, ~99.6% of the N in the atmosphere occurs as 14N. Thus, when an organism obtains N through symbiotic N2 fixation, its own δ 15N value will reflect the relative contribution of N acquired from symbionts compared to N obtained from its diet (i.e., symbiotic N2 fixation will decrease the δ 15N value as it incorporates almost exclusively 14N from the atmosphere). This approach has been employed to estimate the amount of N contributed by microbial symbionts in termites (S48).

This method requires complete knowledge of every food source used by the

insects. In the case of our laboratory maintained leaf-cutter ants, feeding is controlled and involves only two types of leaves. The fraction of N derived from the atmosphere (% Ndfa), through symbiotic N2 fixation, is given the following equation:

% Ndfa = ( [(δ 15Nleaves + ∆dig) − δ 15Nfungus garden] / [(δ 15Nleaves + ∆dig) − ∆fix] ) X 100% where ∆dig (δ 15Nfungus garden − δ 15Nleaves) represents the isotopic discrimination during the digestion of leaf material and ∆fix is the isotopic discrimination resulting from N2 fixation (S49). The δ 15N value for the leaves of five A. platanoides and five Q. ellipsoidalis local trees employed to feed leaf-cutter ant colonies in our lab were 4.61 ‰ ± 0.36 ‰ s.e.m and 3.66 ‰ ± 0.19 ‰ s.e.m, respectively. For δ 15Nleaves we used the mean δ 15N value of both types of leaves (4.13 ‰ ± 0.25 ‰ s.e.m.). We determined the δ 15N value for Atta sp.

13

fungus gardens to be 2.33 ‰ ± 0.16 ‰ s.e.m. (n = 14), thus ∆dig = 1.80. For ∆fix we used the estimated ranges of −2 < ∆fix < 0, as employed by Tayasu and collaborators (S48) for termites. Using this approach, we estimate that the amount of N that the fungus garden obtains from atmospheric N2 fixation in laboratory maintained A. cephalotes colonies ranges from 45% to 61% of its N budget.

In ecological studies a species δ 15N value is used to help determine its sources of N intake as well as the trophic level at which it feeds (S50). This is based on the fact that, as previously mentioned, during catabolic reactions the lighter isotope (14N) is lost at a greater rate, leading to a progressive enrichment of 15N relative to 14N with increasing higher trophic level (S51). In the case of ants, it has been shown that δ 15N is a good predictor of their trophic level when compared with natural feeding observations (S14, S50, S52, S53). Furthermore, isotope-based studies revealed that in the canopies of tropical rainforests many arboreal ants appear to obtain little N through predation and scavenging, suggesting that additional N may be acquired through symbiotic associations with N2-fixing bacteria (S54). Recently it was reported that the mean δ 15N enrichment observed across a single trophic transfer from plants to herbivorous insects is 1.88 ± 0.37 ‰ (S55). In our 15N-enrichment experiments, the fungus garden had an increase in the δ15N value of 1.50 ± 0.12 ‰ (Fig S.5). This increase is within the range of a complete trophic level transfer, further demonstrating that N2-fixers provide an important contribution to the fungus garden N budget.

Even though there are some limitations inherent to 15N-based techniques,

estimates of N2 fixation using stable isotopes are recognized to be more reliable than others, such as those based solely on acetylene reduction. This is because isotope-based measurements reflect N2 fixation integrated over long periods of time (S56). Even though future work refining these estimates is needed, we believe that our initial estimates are fairly accurate, as they are highly congruent with our understanding of N flow in leaves, fungus garden, and exhausted substrate for leaf-cutter ant colonies, both in the field and laboratory. For example, in our study we determined that the N content of the fungus garden is almost double the N content of the leaves (Fig 1C), which fits with the garden obtaining ~50% of its N from N2 fixers. It is possible that the observed higher N content in fungus gardens, compared to leaf input, is solely due to a higher rate of leaf incorporation with respect to removal of exhausted substrate. However, this is unlikely, as there is a relatively constant turnover of the fungus garden both in field colonies (where refuse deposition rates are tightly correlated with harvesting rates (S57)), as well as in laboratory-maintained colonies, which are kept at a constant garden size (their physical space is saturated, preventing significant growth). In addition, our δ 15N-based estimate fit expectations based on several field studies that have analyzed nutrient flow through leaf-cutter ant colonies. In a long-term study on A. colombica in Panama, Wirth and collaborators (S19) found that the N content of exhausted fungus material was only slightly below that of the harvested leaf material, indicating that the colony withdraws only minimal N from the leaves. In contrast, the carbon concentration in the refuse was significantly lower than in the leaf input (S19), suggesting that leaf-cutter ant colonies use leaf material primary as an energy source (see also (S58, S59)). Finally, our estimates

14

are also supported by our finding that the strains of Klebsiella we establish as symbionts of leaf-cutter ant fungus gardens have very high N2-fixing capabilities (see above).

Using our lower estimate for the percentage of N leaf-cutters receive from biological N2 fixation (45%), we performed calculations to evaluate the potential impact for the Neotropical ecosystems in which they occur. Unfortunately, despite leaf-cutter ants being studied for over a century, there is neither a currently accepted estimate for the N budget of mature colonies nor an estimate of the total biomass of fungus gardens within these colonies. Wirth and collaborators (S19) report that each year mature leaf-cutter colonies collect an average of 275 kg dry weight of foraged material, with an N content of 2.4%, and discard approximately 196 kg dry weight (dw) of refused material, with a N content of 2.3%. Therefore, each day, an average of 18 g of N is brought into the gardens from the incoming leaves (275 kg divided by 365 and multiplied by N content) while 12 g of N are discarded (196 kg divided by 365 and multiplied by N content), for a net daily gain of 6 g of N from the harvested leaf material. Based on our isotope-based estimates, these daily N gains from leaves should correspond to 55% of the daily N uptake of the garden, leaving 4.9 g (45%) as the daily amount of N contributed by symbiotic N2-fixing bacteria. Therefore, we estimate that the N2 fixers colonizing the fungus gardens of each mature leaf-cutter ant colony incorporates approximately 1.8 kg of N into terrestrial ecosystems each year. Additionally, given that the fungus garden is turned over every 6 weeks (S60), over a one year period the fungus garden is turned over ~8.7 times. This means that at any given point in time the fungus garden total dry weight is likely greater than 9.1 kg dw (yearly input minus output in terms of foraged material equals a net gain of 79 kg of leaf material incorporated yearly divided by 8.7, the number of times the fungus garden turns-over in a year). Using this information, we estimate that at any given moment, a mature leaf-cutter ant colony contains ~143 g of N acquired directly from the atmosphere by associated N2-fixing bacteria. This is based on the fungus garden having an N content of 3.5 % (Fig. 1C) and receiving ~45% of its N budget from symbiotic N2 fixers (9.1 kg X 0.035 X 0.45 = 0.143 kg).

To compare the importance of symbiotic N2 fixation for the nutritional economy

of different termite species, several authors employ the TDN index (time to double N content, reviewed by Breznak (S61)). We estimated that at any given moment, the fungus garden of Atta sp. colonies contains 319 g of N (9.1 kg x 3.5% N content), and because there are 4.9 g of N fixed daily within these colonies (see above), then N acquired from the atmosphere by symbiotic N2 fixers may support 5.6 N biomass doublings per year. Considering that the fungus garden turns over 8.7 times per year, approximately 60% of each N content doubling could be supported by the symbiotic N2 fixers. Such value falls within our estimate that the symbiotic N2-fixing bacteria can provide as much as 45% to 61% of the fungus garden N budget. It is important to point out that, as opposed to similar studies done with termites, these calculations are based on doubling the biomass of the leaf-cutter ant fungus garden and not the insects themselves. The role of the symbiotic fixers within this system is to contribute to the N economy of the fungus garden, which in turn supports the increase in insect biomass of the colony, and, as discussed above, this additional N fixed from the atmosphere will eventually impact the ecosystems where leaf-cutter ants reside.

15

The contribution of symbiotic N2 fixers to the N economy of insect colonies has also been estimated from acetylene reduction rates. This is the case for classic studies in the well known symbiosis between termites and N2-fixing bacteria. For example, using the acetylene reduction method Prestwich and collaborators (S61, S64) showed that the microbial gut flora of termites of the tropical genus Nasutitermes can fix a significant portion of the nitrogen required by the colony. However, this is not the case for termites in the genus Rhynchotermes, which showed significantly lower acetylene reduction rates, a difference attributed by the authors to the distinctly different diets of the two termite genera (S64). In fact, based on acetylene reduction rates, most termite species appear not to rely on N2-fixing symbionts at all, as reviewed by Breznak (S61), who pointed out that at least some of these extreme variations in AR rates are caused by the N content of the food consumed by termites prior to the test, the age and developmental stage of termites, and variations in life cycle such as bursts of reproductive activity. However, the biggest limitation of the acetylene reduction method is its sensitivity to disturbance of the termite nests, especially physical manipulation and isolation of individual termites from their nest mates and nest material (S65). For example, it has been shown that acetylene reduction activity measured in individual termites declines >90% within 24 hours of nest collection (S62), while a decrease in colony manipulation results in a corresponding increase in acetylene reduction activity (S64). For this reason, and as discussed above, we based our estimates on 15N natural abundance studies, as they reflect the accumulation of fixed N over long periods of time.

We anticipate that contributions of N2 fixers to leaf-cutter ant nutrition likely fluctuate in different stages of their development, at different seasons within the same year (i.e. dry vs. wet season) and in different ecosystems. Due to the tight metabolic regulation of N2 fixation, it is expected that symbiotic microorganisms will fix significantly more N2 when faced with conditions of N starvation or limitation. For example, it is likely that incipient colonies, which have a limited foraging workforce, will have different N requirements than fully mature colonies with millions of ant workers. Also, for leaf-cutter ants inhabiting tropical deciduous forests, N2 fixation could be vital during the dry months, when fresh leaves are not as readily available, as opposed to leaf-cutters that inhabit evergreen rainforests where the supply of fresh plant material is relatively constant throughout the year. Furthermore, prior to their annual reproductive flights, mature leaf-cutter ant colonies produce thousands of virgin queens and males. These alates (reproductive males and females) do not participate in regular worker tasks and create an even higher N need for the colony in order to support the massive production of additional tissue in these large-bodied individuals. Such demands could be met, at least partly, through increased N2 fixation rates. 7. Identification and analysis of nif clusters from N2-fixing bacteria associated with

leaf-cutter ants To further characterize the N2-fixing bacteria associated with leaf-cutter ants, we sequenced the nif clusters from two isolates: Klebsiella variicola At-22 and Pantoea sp. At-9b (See Table S6). Both isolates have fully intact nif clusters spanning a 23kb stretch, with identical gene synteny to the nif cluster of Klebsiella pneumoniae 342, a well-known

16

N2-fixing endophyte (S66). The identified nif clusters encompass a total of 20 open reading frames and include the full-complement of coding regions required for N2 fixation using the classical Molybdenum-Iron (MeFo) nitrogenase pathway (S30). Control of this pathway occurs through the positive and negative regulators nifA and nifL, respectively, both of which are found in this cluster and are known regulators of the nif cluster in other Enteric bacteria (30). To confirm the origin of these two clusters, we calculated the GC content of their respective coding regions and found that they have GC contents of 64.06% and 55.12% for Klebsiella variicola At-22 and Pantoea sp. At-9b, respectively. The GC-content of the nif cluster belonging to K. variicola At-22 is almost identical to the nif cluster in K. pneumoniae 342, which has a GC content of 64.14%. The presence and identification of the nif cluster in these two bacteria associated with leaf-cutter ants also suggests that they are symbionts. Currently, there are 3 other publicly-available sequenced genomes in the genus Klebsiella: the aforementioned K. pneumoniae 342, and the human pathogens K. pneumoniae subsp. pneumoniae MGH78578 and K. pneumoniae NTUH-K2044. Among these, only the plant endophyte K. pneumoniae 342 has the nif cluster, consistent with its role as a symbiotic N2 fixer: it has been reported that this isolate is able to relieve N deficiency symptoms in certain wheat varieties (S67). Furthermore, there is no evidence of the nif cluster in the draft genome sequence of the plant pathogen Pantoea stewartii subsp. stewartii DC283 (Bryan Biehl and Jeremy Glasner, personal communication). Pantoea spp. have also been found to be associated with other plants, including sugarcane and rice, where they also play a mutualistic role as N2 fixers (S68, S69). The identical gene synteny of the nif cluster between K. variicola At-22, Pantoea sp. At-9b, and K. pneumoniae 342 may also suggest how these ant-associated isolates originated. Because leaf-cutter ants in the Neotropics forage on a variety of plant species, there is opportunity for plant-associated Klebsiella sp. and Pantoea sp. to enter the fungus garden and establish themselves as symbiotic N2-fixing bacteria for the ant system (see section 5 above). 8. Detection of genera-specific nifH in the gardens of leaf-cutter ants using real-

time PCR

To verify the presence of genera-specific nifH in the fungus gardens of leaf-cutter ants, we employed a real-time PCR approach. First, we utilized a universal nifH primer set and verified that this gene was present in the fungus gardens of 8 laboratory-maintained Atta cephalotes leaf-cutter ant colonies, as shown in Figure S9a. We then designed primers specific to nifH using all of the sequenced nifH genes from Klebsiella and Pantoea isolates cultured from the fungus gardens of leaf-cutter ants used in this study (Figure S7). Using these primers we test whether nifH from these genera are detectable in leaf-cutter ant fungus gardens (Figure S9b and c). In all 8 colonies, nifH for both Klebsiella and Pantoea were detected in some measure, confirming the presence of leaf-cutter ant-specific nifH for these two genera in the fungus gardens of leaf-cutter ants. We also confirmed that these two primer sets did not cross-react (data not shown). Furthermore, we tested nifH primers for the genus Burkholderia, as it has been reported that this genus can sometimes be isolated from the fungus garden of leaf-cutter ants (S70) and that many species from this genus are known to fix nitrogen in a wide variety of

17

environments (S71). We also found there to be some presence of nifH using this primer set (Figure S9d). These data confirm that nifH is indeed present in the fungus garden, that there are nifH genes specific to leaf-cutter ant-associated Klebsiella and Pantoea, and that there is a community of microbes that may participate in N2 fixation within the ants fungal gardens.

Given that we were only able to culture two Burkholderia isolates from the 80

leaf-cutter ant fungus gardens we tested, and that both of these isolates had insignificant levels of N2 fixation activity measured via acetylene reduction assay (Table S5), it is likely that Burkholderia contributes less fixed N2 than both Klebsiella and Pantoea in the majority of fungus gardens from the leaf-cutter ant systems we studied (see section 2 above). Future work in this area should address the relative contributions of Klebsiella and Pantoea to the overall N economy in leaf-cutter ant fungus gardens, and further if other potential symbionts, including Burkholderia, play a role in N2 fixation.

9. Transmission Electron Microscopy of the Fungus cultivated by leaf-cutter ants.

Recent research demonstrated that some fungi maintain mutualistic associations with N2-fixing endosymbionts in the genus Burkholderia (S72). Given this, and the occasional detection of Burkholderia isolates in the fungus gardens of leaf-cutter ants, we analyzed several garden samples from freshly collected Atta cephalotes colonies by transmission electron microscopy (TEM). Our microscopic analysis confirmed previous descriptions of the fungus cultivated by the leaf-cutters (genus Leucoagaricus), such as multinucleated hyphae without clamp connections, the presence of a dolipore septum characteristic of the Basidiomycota and abundant gongylidia, which are nutrient-rich hyphal swellings upon which the ants feed on (Figure S10). Consistent with previous studies (S73), we observed that fungal hyphae penetrated plant tissue almost exclusively at the cuts made by the ants, thus demonstrating the inability of the fungal cultivar to degrade the plant cell wall by itself. We did not detect, however, any structure resembling a bacterial endosymbiont (i.e. presence of an endogenous microbial cell with an integral membranous system). Similarly, simple stains with DNA-binding fluorescent dyes, such as Hoechst 33342, did not reveal the presence of endosymbiotic bacteria (data not shown). Even though we did not find evidence for the presence of intracellular symbionts in the fungus cultivated by leaf-cutter ants neither within the fungus garden (TEM analysis) nor in pure culture (through nifH detection and AR assays), we cannot conclusively rule out this possibility. Future work addressing this possibility should include more comprehensive microscopic and molecular analyses from a number of different leaf-cutter ant species across different geographic locations and environmental conditions.

18

Supplementary Figures and Tables Figure S1. Acetylene reduction activity for fungus garden samples from three different Atta colonies. Samples from these colonies were analyzed for acetylene reduction activity in independent experiments conducted over a span of two years. Results shown are mean values for three replicates. Error bars represent s.e.m.

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Figure S2. Acetylene reduction activity for different sections of the fungus garden from eight Atta colonies. A. Experiments conducted with laboratory maintained colonies in November, 2005 (n = 3) B. Experiments conducted directly in the field in Costa Rica with five A. cephalotes mature colonies. Leaves correspond to all fragments collected by each colony during a five minute period. Results shown are mean values for three replicates per colony (A) or pooled mean for all colonies evaluated (B). Error bars represent s.e.m.

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Figure S4. Acetylene reduction activity for fungus garden samples for three different Atta colonies under aerobic and anoxic conditions. Acetylene reduction measurements for six fungus garden samples were conducted in the presence of oxygen and under anoxic conditions. In the anoxic treatment, oxygen was exchanged for argon prior to acetylene injection. Results shown are mean values for three replicates. Error bars represent s.e.m.

22

Figure S5. Mean 15N enrichment for different components of A. cephalotes sub-colonies. The mean difference between δ 15N values (N isotopic ratio, per mil [‰]) for control and experimental A. cephalotes sub-colonies is shown. Data correspond to fungus garden, worker ants, and ant brood, one and two weeks after the start of the experiment. Error bars represent s.e.m.

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Figure S6. Preliminary results of 15N2 enrichment experiments show fixed N is incorporated into leaf-cutter ant colonies. Both control and experimental (15N-enriched atmosphere) sub-colonies from A. cephalotes colony MB03-01 survived for a month, functioning normally within their airtight containers. The mean δ 15N value (N isotopic ratio, per mil [‰]) is provided for the different components of these sub-colonies. Results shown were obtained four weeks after the start of the experiment. Error bars indicate s.e.m.

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Figure S7. Bayesian phylogeny based on the nifH gene of confirmed N2 fixers associated with leaf-cutter ants. Isolates of leaf-cutter ant–associated N2 fixers are labeled with the host ant species and color-coded to indicate whether they belong into the genus Klebsiella (blue) or Pantoea (green). The numbers above the branches represent their Bayesian-calculated posterior probabilities.

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Figure S8. Bayesian phylogeny based on the 16S rDNA gene of confirmed N2 fixers associated with leaf-cutter ants. Isolates of leaf-cutter ant–associated N2 fixers are labeled with the host ant species and color-coded to indicate whether they belong into the genus Klebsiella (blue) or Pantoea (green). The numbers above the branches represent their Bayesian-calculated posterior probabilities.

26

Figure S9: Real-time PCR traces for 8 lab-maintained Atta cephalotes leaf-cutter ant fungus garden samples using universal nifH primers (a), Klebsiella-specific nifH primers (b), Pantoea-specific nifH primers (c), and Burkholderia-specific nifH primers (d). For Klebsiella- and Panteoa-specific primer sets, standards efficiencies were determined using pure culture isolates of Klebsiella and Pantoea from leaf-cutter ant fungus gardens, respectively. For Burkholderia, the Klebsiella standard was used because no pure culture isolate from leaf-cutter ant fungus gardens was available. Each trace shows the ∆Rn (normalized net fluorescence signal of PCR product) plotted against the number of PCR cycles.

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Figure S10. Ultrastructure of the fungus cultivated by the leaf-cutter ant Atta cephalotes evaluated by transmission electron microscopy (TEM). A. Gongylidia with central vacuole (scale bar = approx. 1 µm). B. Close up of doliporum septum between two hyphae of the cultivated fungus (scale bar = approx. 1 µm). C. Hyphae seen at lower magnification showing multiple nuclei (scale bar = approx. 3 µm). D. Fungal hyphae between intact plant cells indicating that the cultivated fungus cannot penetrate plant tissue without the cutting action by the ants (scale bar = approx. 3 µm).

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Table S1. Acetylene reduction (AR) activity of fungus garden samples from 35 colonies across eight different leaf-cutter ant species collected in Argentina, Costa Rica and Panama.

1nmol ethylene/hour/g dry weight. Results shown correspond to mean (n = 3) ± s.e.m.

Ant Species Colony code Origin AR fungus garden 1

Acromyrmex echinatior 291 Canal Zone, Panama 0.88 ± 0.10 Acromyrmex echinatior CC031212-01 Canal Zone, Panama 0.55 ± 0.07 Acromyrmex echinatior AP061108-01 Guanacaste, Costa Rica 0.58 ± 0.05 Acromyrmex echinatior AP061104-01 Guanacaste, Costa Rica 0.60 ± 0.02 Acromyrmex echinatior AP061106-01 Guanacaste, Costa Rica 0.72 ± 0.09 Acromyrmex echinatior AP061105-02 Guanacaste, Costa Rica 0.73 ± 0.07 Acromyrmex hispidus-fallax UGM030327-02 Misiones, Argentina 1.19 ± 0.10 Acromyrmex hispidus-fallax SP030327-01 Misiones, Argentina 0.91 ± 0.03 Acromyrmex laticeps UGM030330-05 Misiones, Argentina 0.94 ± 0.05 Acromyrmex laticeps UGM030330-04 Misiones, Argentina 0.54 ± 0.04 Acromyrmex niger UGM030327-03 Misiones, Argentina 1.23 ± 0.09 Acromyrmex niger CC030327-02 Misiones, Argentina 0.90 ± 0.07 Acromyrmex octospinosus CC011020-02 Canal Zone, Panama 0.75 ± 0.23 Acromyrmex octospinosus CC030403-09 Misiones, Argentina 0.86 ± 0.24 Atta cephalotes AL050513-22 Canal Zone, Panama 0.93 ± 0.15 Atta cephalotes BB03-01 Canal Zone, Panama 1.43 ± 0.04 Atta cephalotes CC031212-04 Canal Zone, Panama 1.45 ± 0.17 Atta cephalotes CC031208-10 Canal Zone, Panama 1.60 ± 0.08 Atta cephalotes MB03-01 Canal Zone, Panama 1.64 ± 0.02 Atta cephalotes AP061022-01 Heredia, Costa Rica 0.71 ± 0.06 Atta cephalotes AP061024-05 Heredia, Costa Rica 0.75 ± 0.06 Atta cephalotes CC061021-08 Heredia, Costa Rica 0.76 ± 0.15 Atta cephalotes AP061021-02 Heredia, Costa Rica 0.79 ± 0.08 Atta cephalotes CC061021-03 Heredia, Costa Rica 0.83 ± 0.11 Atta cephalotes AP061026-01 Heredia, Costa Rica 0.93 ± 0.06 Atta cephalotes AP080219-03 Heredia, Costa Rica 1.20 ± 0.10 Atta cephalotes AP080225-02 Heredia, Costa Rica 1.10 ± 0.13 Atta cephalotes AP080304-03 Heredia, Costa Rica 1.22 ± 0.07 Atta cephalotes AP080213-04 Puntarenas, Costa Rica 1.59 ± 0.11 Atta cephalotes RA080212-03 Puntarenas, Costa Rica 0.90 ± 0.07 Atta colombica CC031208-03 Canal Zone, Panama 0.96 ± 0.16 Atta colombica AL031208-02 Canal Zone, Panama 1.04 ± 0.11 Atta colombica CC031208-01 Canal Zone, Panama 1.65 ± 0.08 Atta sexdens CC030329-01 Misiones, Argentina 1.50 ± 0.24 Atta sexdens UGM030330-07 Misiones, Argentina 1.44 ± 0.07

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Table S2: Positive PCR-amplifications of nifH from the mutualistic organisms associated with the leaf-cutter ant–microbe symbiosis.

1 Acetylene reduction on ant workers revealed little nitrogenase activity, suggesting that these successful PCR amplifications likely represent undigested fungus garden material within the ants’ digestive systems.

Ant genus Ant DNA Cultivar DNA Pseudonocardia DNA Fungus Garden DNA

Acromyrmex 0 of 5 0 of 5 0 of 5 3 of 5

Atta 3 of 6 0 of 5 0 of 5 4 of 7

TOTAL 3 of 11 1 0 of 10 0 of 10 7 of 12

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Table S3. Results summary for 15N2-enrichment experiments with eight Atta cephalotes colonies.

1 The experimental (15N2-treated) sub-colony from one of the eight parent colonies collapsed during the second week of the experiment and hence it was removed from the second week analysis. 2 Delta (δ) values (‰), corresponding to each sample’s ratio of 15N to 14N compared to a known standard. 3 Two-tailed paired t-test.

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t-value 3 P-value

Fungus Garden Week 1 8 2.28 ± 0.20 3.19 ± 0.25 0.91 7.45 0.0001

Fungus Garden Week 2 7 1.82 ± 0.34 3.32 ± 0.26 1.50 4.87 0.002

Ants Workers Week 1 8 3.99 ± 0.21 4.47 ± 0.20 0.48 5.28 0.001

Ants workers Week 2 7 3.65 ± 0.22 4.30 ± 0.30 0.65 3.08 0.02

Ant Brood Week 1 8 3.50 ± 0.36 3.80 ± 0.27 0.30 2.47 0.04

Ant Brood Week 2 7 3.40 ± 0.26 3.66 ± 0.11 0.26 1.28 0.247

31

Table S4. δ15N (‰) values for fungus garden, worker ants and ant brood of eight Atta cephalotes colonies employed in 15N2-enrichment experiments after one and two weeks.

1 Values shown correspond to the mean of duplicate or triplicate samples.

2 The experimental (15N2-treated) sub-colony from the parent colony CC061021-08 collapsed during the second week of the experiment and was removed from the second week analysis.

A. cephalotes colony codes Colony component CC031208-10 BB03-01 CC061021-08 AP061022-01 AL050522-01 AP061024-05 CC061021-03 MB03-01

Fungus Garden control 1 2.08 / 2.89 2.54 / 2.75 3.15 / NA 2 2.77 / 2.62 1.71 / 0.90 2.66 / 1.18 1.72 / 0.82 1.63 / 1.58

Fungus Garden 15N 3.38 / 3.05 3.58 / 3.46 3.91 / NA 4.11 / 4.38 2.32 / 2.21 3.28 / 3.37 2.85 / 2.87 2.07 / 3.86

Worker Ants Control 3.25 / 2.80 3.57 / 2.86 3.92 / NA 3.26 / 4.02 4.02 / 3.82 4.68 / 4.35 4.33 / 3.90 4.90 / 3.83

Worker Ants 15N 3.51 / 3.29 3.85 / 3.40 4.45 / NA 4.27 / 4.16 4.54 / 4.25 4.91 / 4.61 4.90 / 4.78 5.32 / 5.60

Ant Brood Control 1.84 / 3.34 2.68 / 3.02 5.18 / NA 4.39 / 4.76 3.55 / 2.44 3.09 / 3.19 3.90 / 3.62 3.36 / 3.41

Ant Brood 15N 2.64 / 3.43 2.83 / 3.72 4.87 / NA 4.66 / 3.90 3.73 / 3.06 3.54 / 3.77 4.11 / 3.86 4.03 / 3.88

32

Table S5. N2 fixation activity measured by acetylene reduction for bacterial isolates obtained in N-free media from fungus garden samples of 80 leaf-cutter ant colonies collected in Argentina, Costa Rica and Panama.

Bacterial isolates 1 Isolates from

Acromyrmex sp. colonies (n = 25)

Isolates from Atta sp.

colonies (n = 55)

Average AR activity 2

Agrobacterium sp. 0 3 0.45 ± 0.26 3

Asaia sp. 2 0 0.10 ± 0.07 Burkholderia sp. 1 1 0.13 ± 0.03 Enterobacter sp. 1 2 0.51 ± 0.23 Klebsiella sp. 4 12 32 190.61 ± 17.78

Pantoea dispersa 1 2 0.27 ± 0.07 Pantoea sp. 4 5 18 21.69 ± 6.35 Rhizobium sp. 0 2 0.69 ± 0.29

Streptomyces sp. 2 0 0.25 ± 0.12 Other genera 5 5 5 0.16 ± 0.04

Azospirillum sp. 6 - - 191.66 ± 31.6 1 Taxonomic identification assigned by sequencing16S rDNA, unless otherwise noted. 2 nmol ethylene/hour. 3 Results shown are the mean activity for all isolates under each category ± s.e.m. 4 Taxonomic identification assigned by sequencing 16S rDNA and 3 other housekeeping genes (gapA, icdA and mtlD). 5 Other genera include single isolates from Cellulosimicrobium, Erwinia, Flavobacterium, Klebsiella oxytoca, Kluyvera, Microbacterium, Nocardiodes, Pseudomonas, Sphingomonas and Stenotrophomonas. 6 Non-ant associated Azospirillum sp. was used as positive control in all tests.

33

Table S6: Topography of the N2 fixation cluster in the draft genomes of Klebsiella variicola At-22 and Pantoea sp. At-9b. The gene, predicted annotation, strand organization, locus coordinates and protein size of each coding sequence in the cluster for both bacteria are shown. Both organisms have identical nif clusters with respect to gene synteny.

Gene Predicted Annotation Strand Klebsiella Locus

Protein Size

Pantoea Locus

Protein Size

nifQ NifQ family protein - 1..516 172 1..507 169

nifB nitrogenase cofactor biosynthesis protein - 516..1922 469 504..1907 468

nifA GAF domain protein/Nif-specific regulatory protein - 2087..3661 525 2072..3643 524

nifL nitrogen fixation negative regulator - 3658..5142 495 3640..5124 495

nifF Flavodoxin + 5462..5992 177 5463..5993 177 mifM nitrogen fixation protein - 6015..6815 267 6064..6849 262 nifZ NifZ family protein - 6815..7258 148 6855..7292 146 nifW putative NifW protein - 7255..7512 86 7289..7546 86 nifV homocitrate synthase - 7515..8657 381 7548..8690 381 nifS cysteine desulfurase - 8673..9875 401 8704..9906 401

nifU Fe-S cluster assembly protein - 9891..10730 280 9936..10760 275

nifX Dinitrogenase iron-

molybdenum cofactor biosynthesis protein

- 10925..11395 157 10964..11371 136

nifN nitrogenase molybdenum-iron cofactor biosynthesis

protein - 11382..12767 462 11409..12782 458

nifE nitrogenase MoFe cofactor biosynthesis protein - 12777..14150 458 12798..14171 458

nifY Dinitrogenase iron-

molybdenum cofactor biosynthesis protein

- 14733..15395 221 14335..14997 221

nifT nitrogen fixation protein - 15406..15624 73 14990..15229 80

nifK nitrogenase molybdenum-iron protein beta chain - 15664..17226 521 15269..16831 521

nifD nitrogenase molybdenum-iron protein alpha chain - 17282..18730 483 16887..18335 483

nifH nitrogenase iron protein - 18747..19628 294 18348..19229 294

nifJ pyruvate

ferredoxin/flavodoxin oxidoreductase

+ 20033..23548 1172 19595..23104 1170

34

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