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Previous Article | Next Article 
Applied and Environmental Microbiology, August 2007, p. 5199-5208, Vol. 73, No. 16
0099-2240/07/$08.00+0 doi:10.1128/AEM.02616-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
,
Jérôme Hamelin,2,
Farma Ndiaye,1
Komi Assigbetse,1
Michel Aragno,2
Jean Luc Chotte,3 and
Alain Brauman3*
Laboratoire d'Ecologie Microbienne des Sols Tropicaux, IRD-ISRA, BP 1386, Dakar, Sénégal,1 Laboratoire de Microbiologie, Université de Neuchâtel, Case Postale 2, CH-2007 Neuchâtel, Switzerland,2 Unité de Recherche SeqBio, IRD, SupAgro, 2 Place Pierre Viala, 34060 Montpellier Cedex 1, France3
Received 9 November 2006/ Accepted 8 June 2007
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ABSTRACT |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The effect of termites on soil is closely linked to their feeding habits and the type of constructions they build (18). Of the six feeding groups described (8), we have chosen to study specifically the microbial compartment of a mound formed by a species of soil-feeding termite (Cubitermes niokoloensis). This choice was based on both the ecological importance of this feeding group (60% of the 2,600 termite species described [11]) and the way in which its mounds are built. This feeding group is the only group to build mounds from a fine mixture of soil and feces containing a dense and specific microbial community. Soil-feeding termite mounds, therefore, provide a useful model for studying the relationship between macrofauna and the soil microbial compartment, which is the principal aim of this study.
Soil-feeding termite mounds (such as those of C. niokoloensis) have very specific properties arising from the combination of materials of two distinct origins, feces and soil (reviewed in reference 5). This creates an increased richness in clay (5 times more), minerals (2 to 3 times more P and Ca and up to 50 times more NH4 [32]), and organic matter (5 to 7 times more C and N [12]) with respect to neighboring soil. The environment for microorganisms derived from soil and feces is modified not just by an increase in available organic compounds but also by a change in their qualities (C/N, humic acid/sugar content [12, 36]) and their availability by the formation of stable clay-humus complexes (14). This richness in organic matter appears to be the reason for the increase in microbial density in termite mounds (3 to 24 times [13]). However, this increase in density is not accompanied by a significant increase in bacterial activity (mineralization) with respect to neighboring soil (31). C. niokoloensis mounds, therefore, provide a site where organic matter is protected from the strong mineralization that is characteristic of the tropical savanna ecosystem. Apart from the physicochemical differences, the termite mounds also have bacterial (13, 17) and fungal (35) communities that are very different from those in neighboring soil.
The aim of this study was to characterize both the structure of the dominant bacterial communities in soil-feeding termite mounds and the specificity of bacterial communities within mounds with respect to the digestive and soil origins of the mound. Using denaturing gradient gel electrophoresis (DGGE) coupled with clonal analysis, the microbial communities in the mound were compared with those of the three main gut segments (anterior gut, midgut, and posterior gut) and with fresh materials used for building mounds (feces and galleries). We were able to demonstrate that the soil-feeding termite mound harbors a bacterial community that differs in terms of structure and diversity from those of its building materials, i.e., termite gut feces and surrounding savanna soil particles, and that the termite mound bacterial community is characterized by the domination of Actinobacteria families. These Actinobacteria clones of soil-feeding termite mound were diverse, distributed among 10 distinct families, and lightly dominated by the Nocardioidaceae family.
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MATERIALS AND METHODS |
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Termite dissection.
Just after the mound was collected in the field, approximately 50 termites were dissected, and the total guts were pooled in 100 µl sodium dodecyl sulfate (4%). This sample was frozen during transport to the laboratory and before DNA extraction. Between 1 and 2 weeks after the mound had been brought to the laboratory, sampling of additional termite guts was completed as follows. Under a hood, individual worker caste termites were dissected in aseptic conditions. By use of two pairs of tweezers, the whole gut of each termite was extracted by pulling the anus with one set of tweezers and securing the head with the other. The whole guts were divided into three segments (Fig. 1): anterior gut, midgut, and posterior gut. About 150 samples of each gut segment were pooled in 2-ml Eppendorf tubes containing 100 µl sodium dodecyl sulfate (4% wt/vol) solution and stored at –20°C before analysis.
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DNA extraction.
Total DNA was extracted from soil samples and from the various gut segments by use of a direct lysis extraction procedure described previously (13). A subsample (0.5 g for soil samples and 0.25 g for gut samples) was prepared using 0.5 g of glass beads (0.1-mm diameter; Biospec Products, Inc., Bartlesville, OK) and 925 µl of sodium dodecyl sulfate buffer (4% wt/vol) in a 2-ml Eppendorf tube (polypropylene; Poly Labo, France). The subsamples were shaken at maximum speed for 5 min using a Biospec 8TM Mini-BeadBeater. After incubation for 1 h at 68°C, 300 µl of 5 M NaCl was added, and the mixture was vortexed and stored for 5 min on ice. This step precipitated the clay to provide optimum DNA recovery. The DNA was precipitated using 40% polyethylene glycol and 2.5 M ammonium acetate successively. The crude DNA was purified using an S400 HR spin column fast DNA purification kit (Pharmacia Amersham, Freiburg, Germany) and then a QIAGEN mini column (QIAGEN, France) using the procedure described by Ranjard et al. (34).
PCR amplification of 16S rRNA gene and DGGE.
PCR amplifications were performed using the forward primer EUB338 (22) with a GC clamp (29) and reverse primer UNIV518 (33). The total reaction mixture (50 µl) contained 2.6 U of Expand Fidelity PCR Taq (Boehringer Mannheim, Mannheim, Germany), 5 µl PCR buffer (10x), 1.5 mM MgCl2, 200 µM of each deoxynucleoside triphosphate, 500 nM of each primer, 0.25 µl of T4 gene 32 protein (Boehringer Mannheim, Mannheim, Germany), sterile water, and about 50 ng of sample DNA. Three replicates were performed for each sample. A Perkin-Elmer GenAmp PCR system 2400 (Perkin-Elmer, Corporation Norwalk, CT) thermocycler was used for PCR amplification with 5 min at 94°C followed by 35 cycles of 1 min at 94°C, 1 min at 55°C, and 1 min at 72°C. The first 20 cycles had an annealing temperature of 65°C, which decreased 0.5°C every second cycle until a touchdown at 55°C. The primer extension was carried out at 72°C for 10 min. The PCR products were checked for size on an agarose gel (1.2% for the DGGE product) stained with ethidium bromide. An 8% polyacrylamide gel was run using a D-Gene system (Bio-Rad Laboratories, Hercules, CA) with a denaturing gradient between 30% and 55%. In addition, one gel including all 10 samples was made so that DGGE patterns for the different compartments could be run under identical electrophoresis conditions. A total of 15 µl of PCR product (mixture of three replicates) was loaded. The gel was run for 5 h at 150 V in Tris-acetate-EDTA buffer (0.5x) at 60°C with a prerun of 10 min at 25 V. After migration, the DGGE profiles were stained with SYBR Green (1:10,000 dilutions) for 20 min and then scanned under UV. The gel images were captured using Bio-capt software (Ets Vilber Lourmat, France).
Gel analysis.
The DGGE band patterns were recovered using Gel Compar software (Applied Maths, Belgium), and the matrices of the relative intensities and distances of migration of the bands were obtained. The various DGGE profiles were compared with the Bray-Curtis distance, and the unweighted-pair group method using average linkages was used for hierarchical cluster analysis to produce the dendrogram by use of R 2.0.1 software (R Development Core Team 2004).
Band extraction and purification.
Because the microbial community profiles of the anterior gut and midgut segments were very similar (about 20%) (see Fig. 2, lanes 1 and 2, respectively; cluster analysis) and the profiles of the posterior gut and whole gut were also very similar (see Fig. 2, lanes 3, 4, and 5; cluster analysis), their respective clone libraries were pooled. A clone library originating from one DGGE profile was set up for each cluster (anterior gut and midgut cluster and posterior gut and whole gut cluster). The DGGE bands were extracted from the gel as described by Jensen et al. (22). These extracts were reamplified and reanalyzed by DGGE to check the electrophoretic mobility. The DGGE bands with the expected mobility were excised from the various samples for sequencing.
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The recombinants were screened by PCR with T7 and SP6 primers to check the size of the inserts, and these PCR products were restricted using HaeIII endonuclease. For each DGGE band, one to three distinct clones with the correct insert size were sequenced.
Construction of 16S rRNA clone library.
A 16S rRNA clone library was constructed from the three compartments: SS, the IW, and the total termite gut. PCR amplifications were performed using the pA (forward) and pH (reverse) (9) primers. The amplification reaction mixture used was the same as that used to amplify DNA for DGGE analysis, as described above. The PCR amplification with a Perkin-Elmer GenAmp PCR system 2400 was as follows: 5 min at 94°C followed by 35 cycles of 1 min at 94°C, 1 min at 55°C, and 1 min at 72°C. The amplicons were purified in 0.8% agarose gel containing ethidium bromide at a concentration of 0.5 µg ml–1 and were recovered from the gel using GFX PCR DNA and gel purification kits (Amersham Biosciences, Germany); then they were cloned into the pCR 2.1 vector (Invitrogen). Competent cells of E. coli DH5
cells were transformed with the ligations, and white colonies were randomly picked and screened directly for inserts by performing colony PCR with M13r and M13f primers.
Sequencing the 16S rRNA clone library.
Plasmid DNA was prepared from the positive clones with the Nucleotrap PCR purification kit (Macherey-Nagel, GmbH, Düren, Germany). Plasmid DNA was then sequenced using an ABI model 3730xl capillary DNA sequencer (Applied Biosystems, Foster City, CA) and a BigDye v3.1 Terminator cycle sequencing kit (Applied Biosystems).
Diversity measurements.
The Shannon diversity index (H') (39) and the Simpson index (D) (40) were calculated according to the following equations:
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Phylogenetic analysis.
All of the sequences were compared with similar sequences from the reference organisms by performing BLAST searches (1). Phylogenetic Actinobacteria trees were constructed with the complete 16S rRNA gene sequences by use of the alignment programs in the ARB package (28), which are based on the rRNA secondary structure. To load our sequences, the nearest neighboring species for each 16S rRNA sequence was found using a BLAST search of the NCBI database. The neighboring sequences were considered to be the references and all were downloaded. The sequences were then aligned using ClustalX. The aligned sequences were imported into the ARB database, with T's already substituted by U's. Consensus-aligned sequences were used to align our sequences with the ARB database. The quick add marked species feature of ARB (parsimony) was used to position our sequences, and the position was manually corrected using the neighboring species found with NCBI BLAST.
Nucleotide sequence accession numbers.
The nucleotide sequence data have been deposited in the GenBank database under accession numbers from AY100703 to AY100745, from AY293289 to AY293299, and from DQ347842 to DQ347944.
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RESULTS |
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Cluster analysis.
A cluster analysis of the DGGE profiles (Fig. 2) was carried out to determine similarities between bacterial communities in the different environments studied. For this analysis, the profiles were categorized into three groups. In the first level, the IW and the SS (lanes 9 and 10) were separated from the other environments (digestive tract, fresh feces, and galleries). Although the bacterial communities of the IW and the SS were clustered in this level, there was less than 18% similarity between these communities. In the next level (about 20% similarity) the bacterial communities in the various segments of the gut (lanes 1, 2, and 3) were separated from the soil reworked by the termites (fresh mound, galleries, and EW). Finally, in the third level (i) the bacterial communities in the gut cluster were split, with the bacterial communities in the anterior gut and midgut (lanes 1 and 2) separated from the communities in the posterior gut and whole gut (lanes 3, 4, and 5), and (ii) the bacterial communities in the reworked soil cluster were split between the freshly reworked soil (fresh feces and galleries; lanes 6 and 7, respectively) and the EW (lane 8).
Sequencing and identification of DGGE fragments.
To determine the affiliation of the dominant bacterial communities in the termite mounds and the other environments, the 46 strongest bands representative of the DGGE profiles were excised, cloned, and sequenced (Table 1). In the anterior gut and midgut (Fig. 2), 14 clones obtained from 11 bands were assigned to five distinct phylogenetic groups (Table 1). Nine clone sequences belonged to the group of Firmicutes. The other groups were spread over four phylogenetic groups.
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In the mound compartments (EW and IW), 7 out of 12 clones (Table 1) originating from 11 excised bands were affiliated with Actinobacteria. To determine whether Actinobacteria species were also present in freshly reworked soil (galleries and feces), six DGGE bands (Fig. 2, lanes 6 and 7) from the lower part of the gel were sequenced. As for the IW, most of the clones (five out of eight) were strongly affiliated to various Actinobacteria families. Of all the clones sequenced, only one (AY293298) was found in both fresh construction material and the IW.
Because most of the bands belonging to the SS profiles were faint, we were able to excise only six dominant bands of the SS samples. Four of the six clones belong to Proteobacteria (Table 1).
Bacterial community structure as determined by clone library analysis.
To complete the DGGE fingerprinting analysis, a 16S rRNA clone library was constructed. A total of 212 clones originating from the mound IW (83 clones), the SS (92 clones), and the termite gut (37 clones) were sequenced (Table 2), using primers that targeted a 650-bp portion of the gene. Rarefaction analysis (Fig. 3) together with coverage estimation (CACE; Table 2) revealed that the sampling depth of the individual clone library was sufficient to cover most of the bacterial diversity of the SS (62%) and, to a lesser extent, that of the termite mound (55%). For the termite gut, the relatively weak coverage estimation (35%) and the slope of the rarefaction curve (Fig. 3) indicated that further sampling may be needed; however, this environment has been already extensively inventoried by previous cloning analysis for a neighboring termite species (38, 41).
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The bacterial compositions at the phylum and phylotype levels differed between the termite gut, mound, and savanna soil (Fig. 4; also see Fig. S1 in the supplemental material). Each compartment was dominated by a particular phylum. The termite mound library was dominated by the Actinobacteria phylum (
50% of all assigned clones), whereas Proteobacteria (mainly the 
subgroup) dominated the SS library (45% of the clones) (Fig. 4). The Actinobacteria phylum represents another important phylum (30% of the clones) in the SS. The termite gut library was dominated by clones affiliated with Firmicutes (Fig. 4; also see Fig. S1 in the supplemental material). The Bacteroidetes/Chlorobi phylum, dominated by bacteria considered to be fermentative, was found as expected in the termite gut library, but interestingly, it was also found to a lesser extent in the mound.
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The 37 phylotypes were classified into 16 different Actinobacteria families (Fig. 5). However, for the termite mound and the SS, there was a clear separation of the clones, depending on their origin. Within the 15 Actinobacteria families, only two families (Nocardioidaceae and Coriobacteriaceae) contained phylotypes from both compartments (Fig. 5). Although the termite gut and the SS did not have any common Actinobacteria phylotypes, nearly all of the Actinobacteria phylotypes (four out of five) from the termite gut were also found in the termite mound.
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For the SS, the 13 phylotypes were distributed in only seven Actinobacteria families, thereby showing a diversity lower than that of the mound environment. Above all, this environment was characterized by the dominance of one family, the Dermabacteraceae, characteristic of the savanna soil, which covered nearly half of all the clones in the SS. The two other important groups were the Geodermatophilus and Micromonospora genera, represented by six and two clones, respectively. The former was not specific to the SS, as it included two clones found in the termite mound.
The five termite gut Actinobacteria clones were affiliated with three families: two specific families, the Coriobacteriaceae (two clones) and the Promicromonosporaeae (one clone), and one nonspecific family, the Nocardioidaceae (two clones), also found in the termite mound.
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DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Shift in the bacterial community structure between the termite gut and the mound.
Because of the dual origin (gut feces and soil particles) of the soil-feeding termite mound, we investigated the bacterial community structures of the various gut sections, feces, mound, and surrounding savanna soil using DNA fingerprinting (DGGE). The cluster analysis of the DGGE profiles (Fig. 2) showed unequivocally that the three compartments investigated (gut, mound, and soil) each harbored different bacterial communities (Fig. 2). There was a major shift in the structure of the bacterial communities between the various segments of the gut and the various mound compartments. This shift is seen in the different distributions of the dominant bands along the DGGE profiles. The strong bands in the gut profiles were situated mainly at the top of the gel (weak denaturing conditions), while the profiles for the mound compartments were all in the lower part of the gel (strong denaturing conditions). This change may indicate that the bacterial community naturally present in the posterior gut is replaced in the mound by a new bacterial community after excretion in the form of feces. This change seems to be very rapid, as fresh feces sampled less than an hour after being deposited in the mound already had a profile that was intermediate between the posterior gut and the mound. This almost rapid change is probably caused by the nature of the environment in the soil-feeding termite gut, which is characterized by several physicochemical (pH, O2, H2) gradients both axially and longitudinally (37). On the other hand, the mound is characterized by a very compact structure rich in organic matter resulting from the fine mixture of feces and soil particles (35). Within the mound, various different bacterial communities coexist in microenvironments (aggregates) created by the termites (13). Passing from an environment that is almost anoxic, alkaline, at micron scale, and half-liquid (gut) to an environment that is oxic, slightly acidic, solid, and rich in available organic matter (mound) may be the cause of the shift observed between the bacterial communities.
The DGGE profiles of the IW is more related (Fig. 2) to the surrounding reference soil than to the mound EW, a mound compartment closer to the fresh termite structures (feces and galleries). This study thus confirmed the differences in terms of bacterial diversity (13) and bacterial activity (32) in termite mounds related to the pedological differences between these two structures (12).
Another interesting finding highlighted by this fingerprinting analysis is the difference between the compositions of the bacterial communities in the reference soil and those in the anterior gut, which should presumably hold soil freshly ingested by the termite. This shows that the anterior gut and midgut have a specific microbial community, even though the physicochemical conditions are not particularly specific (neutral pH and oxic conditions [37]). This study, therefore, confirms the hypothesis, proposed by Schmitt-Wagner et al. (38), that the shift observed from the structure of the digestive bacterial communities to the soil bacterial communities was not caused solely by the extreme pH conditions but also by the lysis of soil bacteria on entry into the gut.
Bacterial composition.
The bacterial community structure in the IW of the mound was strongly dominated (50% of clones) by bacterial sequences affiliated with Actinobacteria families. This result agreed with the DGGE band cloning analysis, where two-thirds of the clones were affiliated with this group. Besides the Actinobacteria, the Firmicutes clones represented another important component of the termite mounds, with nearly 17% of all assigned clones in the library. Nearly half of the mound's Firmicutes clones were from Clostridiales genera closely affiliated with previously retrieved sequences from termite guts (see Fig. S1 in the supplemental material) (38, 41, 42). This indicates that the change in the bacterial community structure between the gut and the mound shown by the DGGE analysis was not complete. The presence of typical anaerobic gut organisms in the mound is not surprising, as the dominance of fine aggregate soil fractions (87% of soil weight [12]) might favor the preservation of conditions for anaerobic and microaerophilic organisms in the mound.
Our investigation of the microbial diversity of soil-feeding termite guts did not completely describe this astonishingly diverse bacterial environment (Table 1), where more than 300 different bacterial phylotypes, representing more than 700 species per termite, have been reported for the gut of the termite Reticulitermes speratus (20, 21). However, our results were in agreement with those obtained by Schmitt-Wagner et al. (38) for a closely related termite species (C. ugandensis) and provided convincing evidence that the bacterial community compositions identified in this study were representative of those of termite gut of this genus (see Fig. S1 in the supplemental material). The gut bacterial community of C. niokoloensis (Senegal) showed that more than 50% of the gut clone libraries were affiliated with the Firmicutes phylum and mostly with the Clostridiales genus (about 50% and 45% of the clones, respectively, by use of DGGE and cloning analysis). Moreover, the majority of the clones were closely affiliated with sequences found also in termite guts (see Fig. S1 in the supplemental material). These results are in good agreement with a study by Hongoh et al. (19), who showed that the majority of termite gut bacteria represent true symbionts intimately linked with termites during their speciation.
A cloning analysis was undertaken from the main DGGE bands and from clone libraries of the main compartments. Comparison of both libraries shows that large proportions (19% and 41% obtained with 97% and 90% of sequence identities, respectively) of DGGE clones (Table 1) were found in 16S rRNA gene clone libraries, indicating a reasonable coverage between the two libraries. The DGGE cloning analysis also confirmed the compartmentalization of the bacterial community in the gut sections investigated. While the Firmicutes, mostly Clostridiales, dominated the anterior gut and midgut communities, the hindgut bacterial community was more diverse, including phyla more characteristic of this part of the gut, such as Spirochaetes, Actinobacteria, and Betaproteobacteria (38). Analysis of the gut clone libraries in this study revealed the presence of Actinobacteria clones (seven clones from all clone libraries), whereas sequences belonging to the Actinobacteria phylum have not been reported in clone libraries from similar termite species like C. orthognathus and C. ugandensis (38). However, the same study (38) reported the detection of Actinobacteria by direct counts using in situ hybridization in the gut. Other reports identified Actinobacteria-related phylotypes specific to the termite gut (belonging to termite clusters) in different termite species (20, 21), including soil feeders (41). Nakajima et al. (30) found that Actinobacteria was one of the dominant phyla colonizing the gut wall of a xylophagous species (R. speratus). Because of their gut wall localization, Actinobacteria species could have been underestimated in these authors' termite gut studies, where they represented less than 1.1% of the total clone libraries in the lumen of the gut (30).
Phylogenetic affiliation of mound Actinobacteria population.
The dominance of the Actinobacteria phylum in the termite mound (Fig. 2) represents the main results of this study. A phylogenetic tree (Fig. 4) clearly demonstrated both the high level of diversity of this Actinobacteria population (41 phylotypes in 16 different Actinobacteria families) and its specificity (70% of families had clones from only one environment).
In spite of its mixed origin, the termite mound had Actinobacteria populations dominated by the Nocardioidaceae family. Only one previous study has reported on the presence of Actinobacteria in soil-feeding termite mounds based on conventional culturing analysis (3). Contrary to our results, the isolates were not significantly different from those in the reference soil, which is not surprising considering that some of the Actinobacteria families (such as Rubrobacteraceae and Coriobacteriaceae) contain phylotypes that have not yet been cultured. Is the dominance of Actinobacteria species due simply to the physicochemical characteristics of the mound being favorable to the development of such a community, or is the community maintained or helped by the termites? Our hypothesis is that C. niokoloensis may cause or help to maintain the dominance of Actinobacteria in its mounds. It has been well established that termites maintain endosymbiotic relationships (digestive bacteria) and exosymbiotic relationships (fungus-growing termites) with microbial communities (4). A symbiotic relationship between insects and Actinobacteria has already been shown in the case of fungus-growing ants (7). This mechanism would imply the reingestion of the IW by the termites as a form of coprophagy, which is widespread among other ecosystem engineers such as earthworms or macroarthropods such as diplopods and isopods (24). Microscopic and ethologic studies are currently in progress to validate this hypothesis.
Although this study was carried out for only one termite mound and its associated population, the results obtained may be considered as representative for this species and even for the genus Cubitermes. Recent studies have shown remarkable stability and uniformity in the bacterial communities of soil-feeding termite mounds of the same species (13) and in different genera of soil feeders (17). This specificity also seems true for fungal populations (35).
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ACKNOWLEDGMENTS |
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We thank Amadou Lamine Ndieng for his technical assistance, two anonymous reviewers for comments that greatly improved the manuscript, and Amy R. Sapkota for proofreading this paper.
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FOOTNOTES |
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Published ahead of print on 15 June 2007.
Supplemental material for this article may be found at http://aem.asm.org/.
Present address: Environmental Microbial Genomics Group, Laboratoire AMPERE UMRCNRS 5005, Ecole Centrale de Lyon, 36 avenue Guy de Collongue, 69134 Ecully Cedex, France.
Present address: INRA, UR 050, Laboratoire de biotechnologie de l'environnement, Avenue des étangs, Narbonne F-11100, France.
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