• Microbial community;
  • Biodiversity;
  • 16S ribosomal RNA;
  • Anaerobic digestion;
  • Archaea;
  • Methanogenesis;
  • Swine manure


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

The microbial community structure of pig manure slurry (PMS) was determined with comparative analysis of 202 bacterial, 44 archaeal and 33 eukaryotic small subunit (SSU) rDNA partial sequences. Based on a criterion of 97% of sequence similarity, the phylogenetic analyses revealed a total of 108, eight and five phylotypes for the Bacteria, Archaea and Eukarya lineages, respectively. Only 36% of the bacterial phylotypes were closely related (97% similarity) to any previously known sequence in databases. The bacterial groups most often represented in terms of phylotype and clone abundance were the Eubacterium (22% of total sequences), the Clostridium (15% of sequences), the Bacillus–Lactobacillus–Streptococcus subdivision (20% of sequences), theMycoplasma and relatives (10% of sequences) and the Flexibacter–Cytophaga–Bacteroides (20% of sequences). The global microbial community structure and phylotype diversity show a close relationship to the pig gastrointestinal tract ecosystem whereas phylotypes from the Acholeplasma–Anaeroplasma and the Clostridium purinolyticum groups appear to be better represented in manure. Archaeal diversity was dominated by three phylotypes clustering with a group of uncultured microorganisms of unknown activity and only distantly related to the Thermoplasmales and relatives. Other Archaea were methanogenic H2/CO2 utilisers. No known acetoclastic Archaea methanogen was found. Eukaryotic diversity was represented by a pluricellular nematode, two Alveolata, a Blastocystis and an Entamoebidae. Manure slurry physico-chemical characteristics were analysed. Possible inhibitory effects of acetate, sulphide and ammonia concentrations on the microbial anaerobic ecosystem are discussed.


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

Intensification of modern pig rearing operations has resulted in greater numbers of animals in large production units that generate large quantities of pig manure slurry (PMS), which is a mixture of pig faeces and urine. In the European Union and in North America, the number of pigs is about 125 and 114 millions, respectively [1–3]. This population generates an estimated pig manure volume of about 100 million tons for the European Union and 91 million tons for North America [4]. Usually, pig manure slurry is stored for a few weeks to a few months in pits located under the pig stalls. Then this waste is commonly stored in outdoor storage tanks for several months prior to discharge into lagoons for treatment and/or spreading on surrounding farmland as fertiliser. While intensive farming methods have proven economically effective, many adverse effects of handling livestock waste have become evident, particularly of slurry [5,6]. The main problems are ammonia volatilisation, offensive odour release (volatile fatty acids, indoles and phenols, ammonia and volatile amines, and volatile sulphur-containing compounds) [7–9], as well as the formation of crust and sediments during storage and the resulting handling difficulties. In addition, other aspects such as the pollution of watercourses via surface runoff and the spread of pathogenic microorganisms are becoming important public issues [10,11].

All these problems are clearly associated, directly or indirectly, to the microbial community composition [12] and to physico-chemical transformations in pig manure slurry [13,14]. Though several studies have been carried out on the anaerobic transformation of the organic matter in manure, only a few studies have investigated the composition of the global microbial community in pig manure slurry and the impact of physico-chemical storage conditions on its evolution [14–19]. Moreover, these studies were often limited by the resolution of culture-dependent microbiological methods, their focus on the fate of pig faecal microorganisms only, or the low number of phylotypes analysed.

During the last decade, molecular techniques have proven very effective for the characterisation of microbial communities in various complex ecosystems, including pig faeces and manure [17–22]. Understanding the processes involved during PMS storage requires a simultaneous analysis of ecosystem parameters and microbial community structure. The present work analyses the pig manure slurry ecosystem in a pig manure storage pit using both physico-chemical analysis of slurry and microbial small-subunit rRNA gene sequence determination.

2Materials and methods

  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

2.1Animals and pig manure slurry sampling

Sampling of pig manure slurry was carried out at an intensive pig rearing farm (SICA Society) located close to Castelnaudary (Aude), in south-central France. The animals, cross-bred from Large White and Landrace and weighing 25–110 kg (fattening pigs), were fed an unchanging cereal formula consisting of wheat and corn (55%), whole wheat (20%), bean forage (10%), soybeans and colza (10%), plus minerals (5%). PMS sampling was carried out at the end of the manure storage period about 60 cm below the surface of a pit containing about 5.4 m3 of slurry resulting from three weeks accumulation of faeces from 60 fattening pigs. About 300 ml were collected anaerobically from three different places in the pit using a pumping system connected to a 500 ml bottle filled with N2 (100%). Samples were homogenised and processed the same day.

2.2Analytical measurements

The pH, redox potential and temperature of the pig manure slurry were measured on site with a glass-electrode pH meter. Redox potential and temperature were measured with a Mettler-Toledo AG model 4300 with a combination of redox electrode (PT48056SC-DPAS-K8S/325). Chemical oxygen demand (COD), total Kjeldahl nitrogen, total ammonia-N and hydrogen sulphide were measured in the raw PMS sample. The soluble fractions of pig manure slurry (supernatants of samples centrifuged and filtered through Nalgene 0.2 μm nylon membrane syringe filters), were analysed for CODs, total carbon (TC), total organic carbon (TOC), inorganic carbon (IC), volatile fatty acids (VFA), soluble Kjeldahl nitrogen, and soluble ammonia-N. All parameters were analysed in triplicate.

The COD was measured using the potassium dichromate–ferrous ammonium sulphate method [23]. TOC content was determined through UV oxidation with a Dohrman DC 90 apparatus. Hydrogen sulphide was determined as described previously [24]. VFA analysis was done on a gas chromatograph fitted with a flame ionisation detector (Chrompac CP 9000) and coupled with an integrator (Chromatopac CR 3A). Total and soluble Kjeldahl-N and ammonia-N were determined by the titrimetric method after distillation using a Büchi 320 apparatus. The TSS (total suspended solids) and VSS (volatile suspended solids) were determined according to standard methods [23].

2.3DNA extraction and PCR amplifications

Total DNA was extracted from 10 ml of homogenised sample as previously described [21]. Bacterial 16S rRNA genes were amplified from the total DNA preparation by PCR amplification using the universal reverse primer w02 (5′-GNTACCTTGTTACGACTT-3′) and the bacterial forward primer w18 (5′-GAGTTTGATCMTGGCTCAG-3′), Escherichia coli positions 1509–1492 and 9–27, respectively [25]. 16S rRNA genes from the Archaea domain were targeted with w02 and the archaeal forward primer w17 (5′-ATTCYGGTTGATCCYGSCRG-3′), E. coli position 3–22. Eukaryotic forward primer W99 (5′-CGGTAATTCCAGCTCC-3′), E. coli position 528–544 [26], and w02 were used for the partial amplification of 18S rRNA eukaryotic genes.

PCRs were as follows: each reaction tube contained 0.2 μg/μl of each primer, 0.2 μg of purified template DNA, 1× REDTaq PCR buffer (Sigma, St. Louis, MO, USA), 2.5 mM (each) deoxynucleotide triphosphate and 1 U of REDTaq DNA polymerase (Sigma), in a total volume of 50 μl. The reaction mixture was prepared on ice and placed in a thermocycler (Perkin Elmer, Forster City, CA, USA) at 94 °C. After an initial denaturation at 94 °C for 2 min, 25 temperature cycles were performed for Bacteria (94 °C for 1 min, 50 °C for 1 min, and 72 °C for 1 min); 30 temperature cycles for Archaea (94 °C for 1 min, 55 °C for 1 min, and 72 °C for 1 min); and 30 temperature cycles for Eukarya (94 °C for 1 min, 49 °C for 1 min, and 72 °C for 1 min). Finally, one step of 72 °C for 10 min was used for all PCRs. The resulting DNA fragments were purified by Qiagen microcolumns according to the manufacturer’ s instructions (Qiagen, Hilden, Germany).

2.4Clone library construction, screening and sequencing

Several clone libraries were made. For each clone library, the purified rDNA fragments from five independent PCR amplifications were pooled together to reduce potential bias. For the bacterial library LD, the cloning was performed using the pGEMt plasmid and E. coli DHI cells as specified by the manufacturer (Promega, Madison, WI, USA). Recombinant cells were selected using ampicillin selection and blue/white screening [27]. The other clone libraries (bacterial LC and LE, archaeal DF and eukaryotic EC) were constructed using the pCR4-TOPO plasmid and TOP10 E. coli competent cells as specified by the manufacturer (Invitrogen, Groningen, The Netherlands). Recombinant cell selection was done by kanamycin resistance and ccd gene killer inactivation.

Archaeal, bacterial and eukaryotic cloned DNA fragments were amplified from positive colonies using plasmid-targeted primers T7 and P13. PCR conditions were: an initial denaturation step at 94 °C for 10 min, 30 temperature cycles of 94 °C for 1 min, 55 °C for 1 min, 72 °C for 1 min and a final elongation step at 72 °C for 10 min. Archaeal and eukaryotic inserts were screened by comparison of Hae III/EcoRI restriction endonuclease cleavage patterns. Eight and five different RFLP patterns were observed for each domain, respectively. Two or more archaeal and eukaryotic inserts from each different restriction pattern were further processed for DNA sequencing. Sequences data confirmed the RFLP screening (data not shown). Otherwise, all bacterial inserts were directly sequenced without RFLP analysis.

DNA sequencing reactions were performed directly on PCR products previously purified on Qiaquick PCR purification kit microcolumns (Qiagen S.A., France). Bacterial and archaeal inserts were sequenced using the “dye-terminator cycle sequencing ready reaction” kit with AmpliTaq DNA polymerase FS kit buffer (Perkin Elmer, Forster City, CA, USA) and primer w31 (TTACCGCGGCTGCTGGCAC), E. coli position 482–500. Eukaryotic inserts were sequenced with the plasmid-targeted primer P13. Reaction sequences were separated using the ABI model 373A sequencer stretch from Applied Biosystems (Perkin Elmer, Forster City, CA, USA). Partial sequences of about 500 bp for Bacteria and Archaea (E. coli position 103–623) and 900 bp for Eukarya (E. coli position 528–1492) were obtained from each insert.

2.5Phylogenetic analysis

Each cloned DNA sequence was compared with sequences available in databases, using Blast from the National Center for Biotechnology Information and the Ribosomal Database Project (RDP-II) [28]. Then all the sequences and their closest relatives were fitted into an alignment of about 12,000 full and partial 16S and 18S rRNA gene sequences using the automated tools of the ARB software package [29]. Partial sequence data were incorporated into trees according to the maximum parsimony criteria without allowing changes to the existing tree topology, using a special tool of the ARB software. The sequences were screened for manual alignment correction and reincorporated into the tree as described above. Distance matrices were calculated to be able to group closely related sequences (97% similarity) within phylotypes. Treeing was performed again from partial sequence data by neighbour-joining method, using Jukes and Cantor correction [30,31]. The stability of tree branches was assessed by the bootstrap method using 1000 replicates. Potential DNA chimeric structure were searched performing fractional treeing on the 5′ and 3′ ends of the sequenced DNA fragments and by analysing the suspected sequences with the CHECK_CHIMERA program (RDP-II) [28].

2.6Nucleotide sequence accession numbers

The nucleotide sequence data reported in this paper appears in GenBank nucleotide sequence database with Accession Nos. AY816753–AY817011.


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

3.1Physico-chemical composition of pig manure slurry

Redox potential, temperature and pH values of PMS were similar to those prevailing in mesophilic anaerobic digestion processes: −358 ± 10 mV, 19.8 ± 0.1 °C and 6.85 ± 0.04, respectively. The PMS contained 128.1 ± 5 g l−1 of total suspended solids composed of about 80% of organic matter (101 g l−1 of volatile suspended solids). Values for total and soluble COD fractions show that this organic matter was essentially particulate (Table 1). Compounds that could be associated with odour production were also high: concentrations of total hydrogen sulphide and VFA were 0.326 ± 0.01 and 16 ± 0.4 g l−1, respectively; 58% of VFA was acetate (9.2 ± 0.2 g l−1). The non-dissociated sulphide concentration, calculated from the raw material measure at pH 6.85 and 20 °C, was 0.198 g l−1[23,32], a level that has been reported to have a significant inhibitory effect on anaerobic digestion [33–35]. Total and soluble ammonia-N were very high and corresponded to 75% and 88% of total and soluble Kjeldahl nitrogen, respectively. However, non-dissociated ammonia calculated at the given temperature (20 °C) and pH (6.85) was low: 0.02 g inline image[36,37]. These data are in agreement with previous reports on PMS characterisation [6,38].

Table 1.  Biochemical characteristics of pig manure slurry
 Soluble fraction (g l−1)Raw material (g l−1)
  1. n.d. = not determined.

  2. aTOC concentration measured is equivalent to 28 g O2 l−1 (COD unit).

Chemical oxygen demand (g O2 l−1)32 ± 0.1115 ± 7.4
Hydrogen sulphiden.d.0.326 ± 0.01
Total Kjeldahl-N4.3 ± 0.59.4 ± 0.9
Ammonia-N3.8 ± 0.67.1 ± 0.7
Total carbon10.9 ± 0.01n.d.
Total organic carbona10.5 ± 0.1 
Total inorganic carbon0.5 ± 0.1 
Volatile fatty acids16 ± 0.4n.d.
Acetate9.2 ± 0.2 
Propionate2.8 ± 0.1 
Butyrate2.6 ± 0.01 
Isobutyrate0.7 ± 0.06 
Isovalerate0.7 ± 0.02 

3.2Molecular inventory of the PMS microbial community

The PMS microbial ecosystem discerned with SSU rRNA gene comparative sequence analysis was extremely diverse in bacterial lineages and remarkably non-diverse in archaeal and eukaryotic lineages. The 202 Bacteria, 44 Archaea and 33 Eukarya sequences analysed were grouped within 108 Bacteria, eight Archaea and five Eukarya phylotypes, on the basis of at least 97% sequence similarity (Table 2). Six chimeric sequences were detected and excluded from the analysis (four Bacteria, one Archaea and one Eukarya).

Table 2.  Phylogenetic affiliation, abundance and diversity of SSU rRNA gene phylotypes from pig manure slurry
PhylotypesaClosest microorganism and/or environmental 16S rRNA gene
NameNo. of clonesb% DivergencePhylotype affiliationcWith more than 97% similarityBetween 97% and 80% similarity% SimilarityIsolated from
  1. n.a. = not applicable; N.D. = not determined.

  2. aClones were sequenced and grouped within phylotypes. The maximum sequence divergence observed within each phylotype is presented.

  3. bNumbers in parenthesis correspond to the number of clones analysed by RFLP (see Section 2).

  4. cThe phylogenetic affiliation of the phylotypes is presented according to the RDP-II. When required, the concordance with Clostridia clustering by Collins et al. [61] is shown in parenthesis. Ru = Rumen uncultured sequence cluster. C.h = Affiliated to Clostrodium hydroxybenzoicum.

Bacteria: 108 phylotypes – 202 clones
Low-GC Gram-positive: 74% phylotypes – 74% clones
Eubacterium and relatives: 19% phylotypes – 22% clones
LC197103C. coccoides (XIVa) C. phytofermentans (AF020431)92Forest soil
LC1830.2C. coccoides (XIVa) C. phytofermentans (AF020431)93Forest soil
LC2430.7C. coccoides (Ru) Clone p-2876-6C5 (AF371584)93Pig gastrointestinal tract
LE9431C. coccoides (XIVa) C. phytofermentans (AF020431)93Forest soil
LD0720.4C. coccoides (XIVa) Clone p-2575-9F5 (AF371606)95Pig gastrointestinal tract
LE6320.7C. coccoides (XIVa)Clone adhufec52 (AF132274) 99Human faeces
LC081 C. coccoides (XIVa)Clone p-189-o5 (AF371642) 99Pig gastrointestinal tract
LC101 C. coccoides (XIVa)Roseburia cecicola (L14676) 98Pig intestine
LC751 C. coccoides (XIVa) Clone BSV51 (AJ229203)93Anoxic rice paddy soil
LE031 C. coccoides (XIVa) C. phytofermentans (AF020431)93Forest soil
LE1031 C. coccoides (XIVa) Eubacterium eligens (L34420)94Pig intestine
LE1091 C. coccoides (XIVa) Ruminococcus gnavus (X94967)94Human faeces
LE271 C. coccoides (XIVa)Ruminococcus obeum (X85101) 99Human faeces
LE361 C. coccoides (Ru) Clone p-2876-6C5 (AF371584)87Pig gastrointestinal tract
LE431 C. coccoides (XIVa)Clone p-2823-6C5 (AF371564) 99Pig gastrointestinal tract
LD4640.4Eubacterium (XI) Anaerovorax odorimutans (AJ251215)89Brackish water sediments
LD7040.4Eubacterium (XI) Clone 3C0d-16 (AB034014)91Rumen
LC761 Eubacterium (XI) Anaerovorax odorimutans (AJ251215)88Brackish water sediments
LE671 Eubacterium (XI) Clone p-3263-42A2 (AF371681)93Pig gastrointestinal tract
LE1001 C. lituseburense (XI) C. glycolicum (X76750)94Mud
LC951 C. propionicum (XIVb) C. lactatifermentans (AY033434)87Caeca of chicken
21 Phylotypes44 Clones      
Clostridium and relatives: 19% phylotypes – 15% clones
LE1222.2C. leptum (IV) Clone p-1877-s962-3 (AF371805)96Pig gastrointestinal tract
LE1721.7C. leptum (IV) Clone p-2215-s959-3 (AF371786)96Pig gastrointestinal tract
LC061 C. leptum (IV) Clone p-953-s962-5 (AF371799)87Pig gastrointestinal tract
LC1881 C. leptum (IV)Clone p-882-a5 (AF371788) 97Pig gastrointestinal tract
LD451 C. leptum (IV) Clone p-953-s962-5 (AF371799)91Pig gastrointestinal tract
LD551 C. leptum (IV)Clone p-900-a5 (AF371807) 97Pig gastrointestinal tract
LE181 C. leptum (Ru) Clone RFN25 (AB009183)92Bovine rumen
LE281 C. leptum (IV)Clone p-1877-s962 (AF371805) 99Pig gastrointestinal tract
LE471 C. leptum (IV) Clone p-953-s962-5 (AF371799)87Pig gastrointestinal tract
LE641 C. leptum (IV)Clone p-2840-6C5 (AF371724) 99Pig gastrointestinal tract
LE911 C. leptum (IV)Clone p-2559-9F5 (AF371715) 99Pig gastrointestinal tract
LE1141 C. leptum (IV) C. viride (X81125)92Human gut/colon
LE1181 C. leptum (IV) Clone UASB brew B86 (AF332721)89Anaerobic digestor
LD3043.2C. botulinum (Ru)Clone P628 (AF261822) 98Manure storage pit
LE6531C. botulinum (I)Sarcina ventriculi (X76649) 98Guinea pig stomach
LD1121.1C. botulinum (I)C. cochlearium (M59093) 98Human faeces
LE9320C. botulinum (I) Clone p-406-o3 (AF371835)91Pig gastrointestinal tract
LE10622C. botulinum (Ru) C. thermocellum (L09173)87Sewage digestor sludge
LD441 C. botulinum (I)Clone p-406-o3 (AF371835) 99Pig gastrointestinal tract
LE891 C. botulinum (Ru) Acetivibrio cellulolyticus (L35516)81Soil
LE1261 C. botulinum (Ru) Clone P628 (AF261822)95Manure storage pit
21 Phylotypes31 Clones      
Clostridium purinolyticum group: 7.5% phylotypes – 5.5% clones
LD3332.2C. purinolyticum (XII)Clone WJGRT-160 (AF175656) 99Contaminated aquifer
LD4120.6C. purinolyticum (XIII) Peptostreptococcus octavius (Y07841)92Human faeces
LD381 C. purinolyticum (XIII)Clone PPC78 (AF445242) 98Pig manure storage pit
LD811 C. purinolyticum (XII) Tissierella praeacuta (X80833)86Soil
LE081 C. purinolyticum (XIII) Helcococcus kunzii (X69837)83Human clinical sources
LE1281 C. purinolyticum (XII) C. hastiforme (X77848)93Soil
LE441 C. purinolyticum (XIII) Helcococcus kunzii (X69837)87Human clinical sources
LE961 C. purinolyticum (C.h) Clone P3IB-23 (AF414571)92Uranium mine sediment
8 Phylotypes11 Clones      
Bacillus–Lactobacillus–Streptococcus: 11% phylotypes – 20% clones
LD40182.2LactobacillusL. reuteri (X76328) 99Human faeces
LC10341.8LactobacillusL. johnsonni (AJ002515) 99Human intestine
LC041 LactobacillusL. amylovorus (M58805) 99Cattle corn silage
LE071 Lactobacillus Pediococcus dextrinicus (D87679)94Fermenting vegetables
LE411 LactobacillusClone p-3424-SwA2 (AF371499) 97Pig gastrointestinal tract
LE841 Lactobacillus Lactobacillus reuteri (X76328)93Human faeces
LC07101.2StreptococcusS. alactolyticus (AF201899) 99Pig intestine
LD341 Bacillus sphaericusClone P610 (AF261818) 98Pig manure storage pit
LE331 Carnobacterium Carnobacterium funditum (S86170)89Antarctic lake water
LE751 Carnobacterium Clone PPC79 (AF445248)96Pig manure storage pit
LC021 EnterococcusClone PPC44 (AF445283) 97Pig manure storage pit
LE391 StaphylococcusJeotgalibacillus psychrophilus (AY028926) 97Fermented seafood
12 Phylotypes41 Clones      
Mycoplasma and relatives: 15% phylotypes – 10% clones
Acholeplasma–Anaeroplasma group
LE3221.8Acholeplasma Acholeplasma axanthum (AF412968)93Pig lung
LE3721.3Acholeplasma Acholeplasma palmae (L33734)92Plant surfaces
LC741 Acholeplasma Mycoplasma feliminutum (U16758)84Oropharynx of cat
LE591 Acholeplasma Mycoplasma feliminutum (U16758)92Oropharynx of cat
LE611 Acholeplasma Mycoplasma feliminutum (U16758)94Oropharynx of cat
LE681 Acholeplasma Acholeplasma axanthum (AF412968)90Pig lung
LE771 Acholeplasma Acholeplasma palmae (L33734)87Plant surfaces
LE791 Acholeplasma Acholeplasma palmae (L33734)81Plant surfaces
LE851 Acholeplasma Mycoplasma feliminutum (U16758)83Oropharynx of cat
LD641 Anaeroplasma Acholeplasma laidlawii (M23932)80Manure
LD391 Anaeroplasma (RCA59)Clone p-1630-c5 (AF371516) 97Pig gastrointestinal tract
LE401 Anaeroplasma (RCA59)Clone p-1630-c5 (AF371516) 98Pig gastrointestinal tract
LD6921.3Clostridium ramosum (XVIII)Catenibacterium mitsuokai (AB030221) 97Human faeces
LD4820.6Eub. cylindroides (XVI)Clone p-2731-24E5 (AF371513) 97Pig gastrointestinal tract
LE351 Eub. cylindroides (XVI)Eub. biforme (M59230) 97Human faeces
LE691 Spiroplasma Clone RF39 (AF001770)91Bovine rumen
16 Phylotypes20 Clones      
Sporomusa and relatives: 2% phylotypes – 1% clones
LE161 Sporomusa (IX)Clone p-922-s962-5 (AF371693) 98Pig gastrointestinal tract
LE1211 Sporomusa (IX)Acidaminococcus fermentans (X77951) 98Pig gut
2 Phylotypes2 Clones      
Flexibacter–Cytophaga–Bacteroides: 17.5% phylotypes – 20% clones
Bacteroides group
LC8221.7Bacteroides fragilisBacteroides sp. 139 (AF319778) 99Pig manure pit
LD191 Bacteroides fragilis Bacteroides uniformis (L16486)82Pig faeces
LE1251 Bacteroides fragilis Clone FPC111 (AF445205)93Pig faeces
LE981 Bacteroides fragilisClone PPC99 (AF445253) 99Pig manure storage pit
LE3841.1Bacteroides splanchnicus Clone BCM3S-11B (AY102902)82Subtropical freshwater marsh
LD54124.7Porphyromonas macacae Clone PPC1 (AF445251)90Pig manure storage pit
LE2431.6Porph. macacae Bacteroides merdae (X83954)85Human faeces
LC19320.2Porph. macacaeClone PPC1 (AF445251) 99Pig manure storage pit
LD5320.5Porph. macacae Clone RFN47 (AB009203)93Bovine rumen
LC171 Porph. macacae Clone RFN41 (AB009197)90Bovine rumen
LD171 Porph. macacae Clone p-987-s962-5 (AF371910)95Pig gastrointestinal tract
LD871 Porph. macacae Bacteroides merdae (X83954)91Human feces
LC9122.3Prevotella nigrescens Clone p-2443-18B5 (AF371893)95Pig gastrointestinal tract
LE0521.5Prevotella nigrescensClone p-2517-18B5 (AF371893) 97Pig gastrointestinal tract
LE451 Prevotella nigrescensClone p-993-s962-5 (AF371898) 99Pig gastrointestinal tract
LE151 Prevotella nigrescens Clone p-3951-23G5 (AF371897)88Pig gastrointestinal tract
Cytophaga group I
LD6320.1Flavobacterium Flavobacterium breve (M59052)92Human intestine
LC921 Flavobacterium Clone Bihii40 (AJ318144)91Industrial gas biofilter
LE951 N.D. Clone p-2202-s959-3 (AF371919)92Pig gastrointestinal tract
19 Phylotypes41 Clones      
Minor phyla: 8% phylotypes – 6% clones
Spirochetes and relatives: 2.5% phylotypes – 2.5% clones
LD5232.7Spirochaeta halophila Clone IIIB-9 (AJ488100)92Chlorine removing sludge
LC961 Spirochaeta halophila Wall less Spirochaeta sp. (M87055)91Kusaya gravy
LD061 Treponema Treponema brennaborense (Y16568)86Cow digital dermatitis
Proteobacteria group: 3.5% phylotypes – 2.5% clones
LE2920.2P. gamma, Xanthomonas Schineria larvae (AJ252146)93Wohlfahrtia magnifica larvae
LE821 P. gamma, PseudomonasP. halodenitrificans (X90867) 98Groundwater underlying a pig rearing facility
LD711 P. delta Clone p-4193-6Wa5 (AF371949)90Pig gastrointestinal tract
LE1101 P. alpha, Rhodobacter Paracoccus denitrificans (X69159)95Soil
Others: 2% phylotypes – 1% clones
LC1011 Flexistipes sinusarabici assemblage Clone W028 (AF125202)82Human oral cavity
LC1021 Fibrobacter group Fibrobacter intestinalis (M62695)88Rumen
9 Phylotypes12 Clones      
DF86273.9CA11 groupClone Ar28 (AF157523) 98Pig manure storage pit
DF1620.5CA11 groupClone Ar21 (AF157522) 99Pig manure storage pit
DF5320.5CA11 groupClone Ar26 (AF157521) 98Pig manure storage pit
DF7261MethanobacterialesMethanobrevibacter smithii (U55233) 99Intestinal tract of animals
DF1032.9Methanobacteriales Methanosphaera stadtmanii (M59139)95Intestinal tract of animals
DF0221Methanomicrobiales Methanofollis liminatans (AF095271)94Intestinal tract of animals
DF191 MethanomicrobialesMethanoculleus olentangyi (AF095270) 99Intestinal tract of animals
DF331 Methanomicrobiales Methanogenium organophilum (M59131)96Intestinal tract of animals
8 Phylotypes44 Clones      
EC0518 + (9)n.a.Nematoda/RhabditidaRhabditella axei (U13934) 99Soil
EC081 + (1)n.a.BlastocystisBlastocystis sp. SY94-7 (AB091249) 97Pig and human intestines
EC121 + (1)n.a.Alveolata Clone BOLA176 (AF372786)86Lagoon sediments
EC251 Entamoebidae Entamoeba moshkovskii (AF149906)88Polluted water and sediments
EC541 Alveolata Clone BOLA176 (AF372786)89Lagoon sediments
5 Phylotypes33 Clones      

3.3Bacterial diversity

The majority of the sequences recovered from the pig manure slurry were not closely related to sequences in public databases. From the 108 bacterial phylotypes, only 39 (36%) presented more than 97% similarity with previously known sequences. Forty-four (41%) showed similarity values in the range of 97–90%; the others (23%) had similarity values between 90% and 80%. Only 13% of all phylotypes were identified as known and cultured species, all but two belonging to the low-GC Gram-positive bacteria. This phylogenetic group contained 74% of the phylotypes observed and 74% of the bacterial sequences analysed. Other phylotypes belonged to the Flexibacter–Cytophaga–Bacteriodes (20% of bacterial sequences), the Spirochetes and relatives (2.5%), the Proteobacteria (2.5%), the Fibrobacter group (0.5%) and the Flexistipes sinusarabici assemblage (0.5%).

The four most abundant phylotypes were related to: Lactobacillus reuteri (X76328) (LD40, 9% of the bacterial sequences); a pig manure clone loosely related to Bacteroides forsythus (LD54, 6% of sequences); Steptococcus alactolyticus (LC07, 5% of sequences) and Clostridium phytofermentans (LC197, 5% of sequences). Thirty-one phylotypes (29%) contained between four and two sequences and 73 phylotypes (68%) contained only one sequence. The estimated percent coverage of the 202 analyzed bacterial sequences is about 46%[39].

The Eubacterium and relatives (RDP Reg. No. 2.30.4), the most abundant and diverse bacterial group found in the inventory, included 22% of the total bacterial sequences and 21 phylotypes (Table 2). Only five phylotypes of this group had more than 97% sequence similarity with known sequences from databases; two of them, LC10 and LE27, were identified as Roseburia cecicola, isolated from pig intestine, and Ruminococcus obeum, isolated from human faeces, respectively. The majority of the Eubacterium and relatives phylotypes (15 out of 21) belonged to the Clostridium coccoides (cluster XIVa) subgroup (RDP registration (Reg.] No. Numerically dominant phylotype LC197 was only distantly related to the forest soil bacterium Clostridium phytofermentans and clustered with four other phylotypes (LC18, LE03, LE94, and LC75) (88% similarity in the cluster but a low bootstrap value of 40%). Taken together, they contained up to 9% of the bacterial clones. All the other phylotypes were related to sequences or microorganisms retrieved from pig or human faeces. Six other phylotypes from the Eubacterium and relatives group belonged to the Eubacterium (Clostridium cluster XI), the C. lituseburense (cluster XI) or the C. propionicum (cluster XIVb) subgroups (Table 2).

A total of 29 phylotypes were affiliated to Clostridium groups. Eight of them belonged to the C. purinolyticum group (clusters XII and XIII or RDP Reg. No. 2.30.5) and contained about 5.5% of the bacterial sequences analysed. They were not closely related to any known sequence, except for phylotypes LD33 and LD38 that presented more than 98% similarity with clones from a livestock-contaminated aquifer and a pig manure pit (Table 2). The 21 other Clostridium phylotypes (15% of bacterial sequences) were split between the C. botulinum (cluster I or RDP Reg. No. and C. leptum subgroups (cluster IV or RDP Reg. No. (Table 2). The most striking observation was the strong similarity between 10 of the 13 phylotypes from the C. leptum subgroup (RDP Reg. No. and phylotypes originating from the pig gastrointestinal tract molecular inventory published by Leser et al. [22]. Three of them (LD55, LE12 and LE28) grouped around Eubacterium desmolans isolated from cat faeces (85% similarity in the cluster and bootstrap value of 60%) while two others (LE64 and LE91) grouped around Faecalibacterium prausnitzii isolated from other animal faeces (90% similarity in the cluster and 73% bootstrap value). Within the C. botulinum subgroup, phylotypes LE65 and LD11 were identified as Sarcina ventriculi and C. cochlearium isolated from a guinea pig stomach and human faeces, respectively. Phylotypes LD30 and LE126 clustered with a pig manure pit uncultivated bacterial clone (95% similarity in the cluster, bootstrap value of 96%).

The 12 phylotypes (20% of the bacterial clones) affiliated with the Bacillus–Lactobacillus–Streptococcus group (RDP Reg. No. 2.30.7) presented two interesting features. Nine of them had more than 96% similarity to previously known sequences or species. It is the highest rate observed for any of the analysed groups (Table 2). Two of them (LD40 and LC07) were numerically most abundant phylotypes of the inventory (Table 2). LD40, identified as Lactobacillus reuteri, contained 18 sequences, which represents 9% of the bacterial sequences. LC07 (10 sequences, 5% bacterial sequences) was the only phylotype affiliated with the Streptococci (RDP Reg. No. Its closest relative was Streptococcus alactolyticus (99% similarity).

Sixteen phylotypes (10% of bacterial sequences) were affiliated with the Mycoplasma and relatives (RDP Reg. No. 2.30.8) (Table 2). All but one belonged to the Acholeplasma–Anaeroplasma group (RDP Reg. No. Only five out of the 15 phylotypes had more than 97% sequence similarity with known sequences. The other nine phylotypes were only loosely related to cultured species, with a mean sequence similarity of 87.6% to their closest relative. They clustered around Mycoplasma feliminutum, of feline origin (82% similarity in the cluster, four phylotypes); Acholeplasma palmae isolated from plant surfaces (79% similarity, three phylotypes); and A. axanthum isolated from pig lung (84% similarity, two phylotypes) (Fig. 1). The LE69 phylotype was affiliated with a Spiroplasma group (RDP Reg. No. containing, also, two uncultured bacterium sequences with rumen and pig gastrointestinal tract origins.


Figure 1. Evolutionary distance tree of the Mycoplasma-related sequences (RDP Reg. No. 2.27). Tree was constructed using the ARB software package. The out-group sequence used in the analysis was Thermotoga maritima (M21774). Number of sequences in PMS phylotypes is indicated in brackets. Bootstrap values higher than 50% are shown in the tree.

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Only two phylotypes (1% of total sequences) were affiliated with the Sporomusa and relatives (RDP Reg. No. 2.30.3). They presented 98% sequence similarity with a pig gastrointestinal tract sequence (LE121) and with Acidaminococcus fermentans isolated from pig gut (LE16) (Table 2).

The Flexibacter–Cytophaga–Bacteroides group (RDP Reg. No. 2.15) was represented by 19 phylotypes (20% of total sequences) (Table 2). Most of them (16 phylotypes) belonged to the Bacteroides group (RDP Reg. No. and had, for their closest relative, sequences retrieved from pig faeces or manure storage pit. Only one phylotype was identified as a cultured species: the sequence LC82 had 99% similarity with the sequence from Bacteroides sp. 139 isolated from a pig manure pit. The second numerically most abundant phylotype of the inventory (LD54, 12 sequences) clustered with the LC193 phylotype (two sequences) and an uncultured sequence isolated from pig manure within the P. macacae subgroup (90% similarity in the cluster and 97% bootstrap value). Finally, three phylotypes were classified outside the Bacteroides group. Two were somewhat related (around 90% similarity) to Flavobacterium sequences (LD63 and LC92). The last one, phylotype LE95, could not be precisely assigned to any subgroup. However, this phylotype clustered with a pig gastrointestinal tract sequence with a bootstrap value of 94%.

The last nine phylotypes were spread within phylogenetic groups that were only poorly represented within the molecular inventory. Three phylotypes were affiliated with the Spirochetes and relatives (RDP Reg. No. 2.27) with bootstrap values higher than 94%. However, their closest relatives had a similarity value lower than 92% and were of diverse origins (Table 2). None of them was related to sequences originating from swine or manure or any digestive tract. Four phylotypes were affiliated to the Proteobacteria groups (RDP Reg. No. 2.28) (Table 2). Two phylotypes belonged to the Gammaproteobacteria, LE82 being identified as Pseudomonas halodenitrificans (98% sequence similarity), a heterotrophic denitrifier isolated from groundwater underlying a pig rearing facility. Another belonged to the Alphaproteobacteria and was related to the heterotrophic denitrifierParacoccus denitrificans (LE110, 95% sequence similarity). Finally, one (LD71) belonged to the Deltaproteobacteria and was only slightly related to an uncultivated PGT clone (90% sequence similarity) but not strongly linked to any subgroup. Finally, LC102 was only distantly related to Fibrobacter intestinalis (88% sequence similarity and bootstrap value of 99%) from the Fibrobacter group (RDP Reg. No. 2.25) (Table 2). In the Flexistipes sinusarabiciassemblage (RDP Reg. No. 2.14), LC101 was related to an uncultured bacterial clone W028 isolated from a human oral cavity (82% sequence similarity and bootstrap value of 92%).

3.4Archaeal diversity

Archaeal diversity was limited to only eight phylotypes that were all closely related (more than 94% similarity) to already known sequences retrieved from manure or the intestinal tract of animals (Table 2). The estimated percent coverage of the library by the archaeal sequences analysed was 82%[39].

The DF86 phylotype (27 sequences) strongly dominated the library and contained 61% of the total archaeal sequences. This phylotype, as well as two others (DF16, DF53) was closely related to uncultured clones isolated from a pig manure storage pit (about 98% sequence similarity) [17]. They all belonged to a phylogenetic group exclusively composed of sequences from uncultured microorganisms from diverse anaerobic environments (Fig. 2): anaerobic digester processes [21,40], contaminated aquifer [41], rumen fluid [42], insect hindguts [43]. Their closest, though still very distantly related, cultured species (less than 77% sequence similarity) is the non-methanogenic Thermoplasma acidophilum.


Figure 2. Evolutionary distance tree of the “CA11”Archaea-related sequences. Tree data and symbols are as in Fig. 1.

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The other five archaeal phylotypes were affiliated with methanogenic H2/CO2 utilisers (Table 2) and belonged to the Methanobacteriales group (DF72 and DF10; RDP Reg. No. 1.1.2) or the Methanomicrobiales group (DF19, DF02 and DF33; RDP Reg. No.

3.5Eukaryotic diversity

The analysis of the 33 eukaryotic sequences showed a low diversity of only five phylotypes (Table 2). The most abundant phylotype (EC05, 27 sequences) was related to the free-living soil saprobe nematode Rhabditella axei with a sequence similarity of 99%. The presence of this pluricellular eukaryote, represented by 80% of the eukaryotic sequences, caused a bias in the results observed for abundance and diversity in this library. Only a few other eukaryotic phylotypes were found. One phylotype (EC08) was related to the Blastocystis sp. SY94-7 (97% sequence similarity). This protozoan has been isolated from faecal samples of pigs and zoo animals [44,45].

The last three phylotypes were only moderately related to an uncultured alveolate isolated from lagoon sediments (EC12 and EC54; 86% and 89% sequence similarities, respectively); or to Entamoeba moshkovskii a free-living amoeba isolated worldwide from sewage, as well as from river and lake sediments (EC25, 88% similarity) [46].


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

The pig manure slurry microbial ecosystem discerned with small subunit rRNA gene cloning and analysis proved to be very diverse in the Bacteria lineage while it was not diverse in the Archaea and Eukarya lineages. The Bacteria phylotypes belonged mainly to the usual fermentative microbial groups found in anaerobic digestion ecosystems: the low-GC Gram-positive bacteria (Eubacterium, Clostridium, Bacillus–Lactobacillus–Streptococcus) and the Bacteriodes. This bacterial community structure is very similar to the one obtained by Leser et al. [22] in pig gastrointestinal tract (PGT) (Fig. 3). Indeed, 33% of PMS phylotypes were closely related to phylotypes retrieved from pig gastrointestinal tract, many of them within the C. leptum (cluster IV) subgroup and the Bacteroides (Table 2). This similarity, which was expected given that manure slurry is directly seeded by pig faeces, was accentuated by the fact that S. alactolyticus and L. reuteri belonged to the most abundant phylotypes in both ecosystems. Nevertheless, several differences could be observed between both ecosystems. At the group level, pig manure slurry contained higher number of sequences affiliated to the C. purinolyticum (clusters XII and XIII), the Mycoplasma and relatives and the Flexibacter–Cytophaga–Bacteroides groups (5%, 10% and 20% of sequences, respectively) than the pig gastrointestinal tract (0.05%, 3% and 9% of sequences, respectively) [22]. Closer comparison of each phylogenetic group revealed several highly represented phylotypes in pig gastrointestinal tract that were clearly under-represented within pig manure slurry. This was the case for the phylotypes related to: (i) Faecalibacterium prausnitzii (about 10% of sequences in PGT and 0.5% in PMS) and Sporobacter termitidis (about 6% of sequences in PGT and none in PMS) from the Clostridium and relatives group and (ii) for Eubacterium rectale ATCC 33656 (T) (about 5.7% of sequences in PGT and none in PMS) from the Eubacterium and relatives group. Another difference was observed within the Mycoplasma and relatives, where phylotypes from PGT were exclusively affiliated to the Eubacterium cylindroides subgroup (about 2.5% of sequences in PGT) (Clostridium cluster XVI or RDP Reg. No. while they belonged in majority to the Acholeplasma–Anaeroplasma group in PMS and represented about 7% of the total sequences. These observations are in agreement with data from all other molecular inventories published on pig gastrointestinal tract and pig manure slurry [18,22,47,48].


Figure 3. Comparative abundance and diversity of major bacterial lineages found in pig manure slurry (▪, black bars) and pig gastrointestinal tract (□, empty bars). Roman numbers refer to Clostridium clustering by Collins et al. [61]. Ru., Rumen uncultured clusters; BLS, Bacillus–Lactobacillus–Streptococcus; FCB, Flexibacter–Cytophaga–Bacteroides.

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In contrast to that of Bacteria, the Eukarya library revealed only five phylotypes. However, the presence of a pluricellular nematode may have prevented the observation of an underlying diversity. Three phylotypes were not closely related to previously known eukaryotic sequences, a fact which underlines the need to explore the phylogeny of these organisms more deeply. Their role in pig manure slurry is unknown. One phylotype was affiliated to Blastocystis sp. This parasite has been detected previously by light microscopy in pig faeces and pigs with diarrhoea but no correlation was found between intensity of Blastocystis infection and the occurrence of diarrhoea [44,45].

Finally, low diversity was observed in the Archaea in pig manure slurry as only eight phylotypes were found. Five of them were identified as previously known cultured hydrogen-utilizing methanogenic Archaea. These species are usually present in the intestinal tract of animals when methane production via H2 becomes significant [49,50]. The three other phylotypes (representing 70% of the total Archaea sequences) clustered with the dominant phylotypes from another pig manure molecular inventory [17] and belonged to a group of sequences, retrieved exclusively from uncultured microorganisms from diverse anaerobic habitats. From distance matrix calculation, all sequences from this group presented a minimum of 90% similarity between each other (data not shown). This cluster was named CA11 in accordance with its first published sequence [21]. Since the cluster shows no evident relationship to any archaeal group, the metabolic activity of the corresponding microorganisms remains unknown. Their closely related cultured species (less than 77% sequence similarity) belong to the aerobic thermophilic acidophilic non-methanogenic Thermoplasmales group. No phylotypes related to known acetoclastic methanogens were observed in the inventory.

The big difference in diversity between the Bacteria and Archaea domains found in pig manure slurry has also been observed in various anaerobic digestion ecosystems, such as an anaerobic digester [21], termite intestinal tract [51,52], rumen [44,53], a contaminated aquifer [41] and sediments [54,55]. This is due to the simplification of substrates, both in number and complexity, through the trophic chain involved in anaerobic degradation of organic matter [56]. However, the absence of known acetotrophic methanogens is surprising: such an absence is typical of animal digestive tracts where methane production from acetate is reduced because of the absorption of volatile fatty acids through the animal intestinal epithelium [50,57]. This observation may be explained by different hypotheses: the “CA11”-related phylotypes might actually form a group of as yet uncharacterised acetotrophic methanogens. However, this hypothesis is not supported by the observations that acetate accumulates in manure and that these phylotypes where also dominant within pig faeces (data not shown). An alternative hypothesis might be that since the microbial community present in pig gastrointestinal tract provides the main seeding of PMS, the amount of acetotrophic methanogens coming through to the manure may be very low and their establishment at a detectable level may require a longer period than three weeks (time of manure sampling in this study). Moreover, the increasing concentrations of VFA and non-dissociated sulphide observed during PMS storage may further inhibit their development [33–35,58]. Such an absence of effective acetotrophic methanogens in the seeding of pig manure slurry, combined with the rapid inhibitory effects of environmental parameters, may break the syntrophic association required for acetate degradation in the manure slurry [59,60]. It would result in the absence of methane production from acetate with the resulting accumulation of acetate, which is effectively the case in pig manure slurry.


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References

This work was supported by the GIS (Group of Scientific Interest) Porcherie Verte. We thank the Mexican Ministry of Science and Technology (CONACyT) for the award of a scholarship for R.S-C.


  1. Top of page
  2. Abstract
  3. 1Introduction
  4. 2Materials and methods
  5. 3Results
  6. 4Discussion
  7. Acknowledgements
  8. References
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