Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Effect of Storage Period on the Changes of Odorous Compound Concentrations and Bacterial Ecology for Identifying the Cause of Odor Production from Pig Slurry

  • Ok Hwa Hwang,

    Affiliation National Institute of Animal Science, Rural Development Administration, Wanju-Gun, Jeollabuk-Do, Republic of Korea

  • Sung Back Cho,

    Affiliation National Institute of Animal Science, Rural Development Administration, Wanju-Gun, Jeollabuk-Do, Republic of Korea

  • Deug Woo Han,

    Affiliation National Institute of Animal Science, Rural Development Administration, Wanju-Gun, Jeollabuk-Do, Republic of Korea

  • Sang Ryoung Lee,

    Affiliation National Institute of Animal Science, Rural Development Administration, Wanju-Gun, Jeollabuk-Do, Republic of Korea

  • Jeong Hoon Kwag,

    Affiliation National Institute of Animal Science, Rural Development Administration, Wanju-Gun, Jeollabuk-Do, Republic of Korea

  • Sung Kwon Park

    Affiliation Department of Food Science and Technology, Sejong University, Seoul, Republic of Korea

Effect of Storage Period on the Changes of Odorous Compound Concentrations and Bacterial Ecology for Identifying the Cause of Odor Production from Pig Slurry

  • Ok Hwa Hwang, 
  • Sung Back Cho, 
  • Deug Woo Han, 
  • Sang Ryoung Lee, 
  • Jeong Hoon Kwag, 
  • Sung Kwon Park


Odor from buildings where pigs are housed is generated by anaerobic fermentation of undigested materials in pig slurry stored for several weeks in pit. The objective of this study was to investigate the effect of storage period on the level of odorous compounds in pig slurry and on its bacterial community. A slurry sample (15 L) was taken from the pit of a finisher pig building and incubated in acryl chambers for six- weeks. Slurry for analysis was sampled every two-week. Levels of odorous compounds in the slurry sample were drastically changed after two weeks of storage period; levels of phenols and short chain fatty acids (SCFAs) were decreased (P<0.05), whereas indoles and branched-chain fatty acids (BCFAs) were increased (P<0.05). Among dominant bacteria, Bacteroides and Porphyromonadacese_uc_g revealed a strong positive correlation with the levels of phenols and SCFAs. Populations of AC160630_g, Acholeplasmatales_uc_g, Mollicutes_uc_g and Cloacamonas_f_uc_g positively correlated with indole and BCFAs content. Taken together, levels of odorous compounds were increased after two weeks of storage, possibly because of changes in the predominant bacterial groups to those that use protein as a carbon source in the hypo-carbohydrate conditions.


Large amounts of pig slurry are generated by intensive animal farming and industrial livestock production (factory farming); in South Korea, the amount increased from 4,370 million tons in 2009 to 4,724 million tons in 2013 [1]. Swine farming accounts for more than 54% of civil complaints about odor from livestock facilities in South Korea [2]. Odor emissions from pig farms are mainly affected by the type of pig house; in South Korea, 76% of pig houses are built with an open ventilation system and 52% with a slurry storage system [2]. To cut down livestock odor, the Korean government enacted an offensive odor control law [3] and restricted the zones where livestock buildings are constructed [4]. Odor control is a prerequisite for successful and sustainable pig production coexisting with nearby residents [5].

Pig slurry is usually stored for a couple of weeks to months inside a pit under the pig building before being cleaned out. During this storage period, anaerobic fermentation triggered by microbes using the undigested nutrients and endogenous materials in the slurry is the main cause of malodor generation [6]. Surface crust and sediment layer formation is another cause of odor [7]. More than 200 chemical compounds are known to be related to livestock odor, but only several compounds are contributed to the offensive odor [810]. Recently, researchers identified amines, ammonia, volatile fatty acids (VFAs), phenols and indoles as key odorants in feedlot manure [5, 914].

Most of the odor-causing materials are produced by protein degradation. If carbohydrates are scarce in pig slurry during the storage period, protein becomes a primary source of fermentable carbon [15]. The nutritional composition of slurry also modulates the bacterial community and its end-metabolites [16]. A few studies have investigated the causes of odor, indicating that there is a correlation between the composition of the bacterial community and odorous compounds [17, 18]. However, these correlations are hard to interpret because of the rapidly changing environment of pig production, including feed ingredients and the physical condition of the pigs.

The objectives of this study were to identify the cause of odor from pig houses with regard to the effect of the slurry storage period on the changes in concentration of odorous compounds and the composition of bacterial communities.

Materials and Methods

Experimental design and slurry collection

Fresh slurry was collected from the pit under a pen housing finisher pigs [total of 60 {(Landrace × Yorkshire) × Duroc}] with body weight (BW) of 80~110 kg in National Institute of Animal Science, Wanju-Gun, Jeollabuk-Do (GPS: 35°49´26.7"N, 127°03´31.6"E). The finisher pigs were fed a basal diet formulated according to the Korean Feeding Standard [19]. Fifteen liters of slurry were incubated in an acryl chamber for 6 weeks at room temperature (20~25°C). Air was continuously supplied to the upper compartment of the chamber with 15 mL/min velocity [20]. Slurry was sampled every 2 weeks for analysis of odorous compounds and the bacterial community. Odorous compounds including VFAs [short chain fatty acids (SCFAs) and branched-chain fatty acids (BCFAs)], phenols and indoles were measured by gas chromatography (GC). The bacterial community was analyzed by pyrosequencing using the 454 FLX Titanium System (Roche, Pleasanton, CA, USA).

Odorous compound analysis

Volatile Fatty Acids.

Five milliliters of slurry were mixed with 1 mL of 25% meta-phosphoric acid solution (Sigma-Aldrich, St. Louis, MO, USA) and 0.05 mL of saturated mercury (II) chloride solution (Sigma-Aldrich, St. Louis, MO, USA) in a 15 mL plastic tube. The mixed solution was then centrifuged at 3,134 × ɡ for 20 min at 20°C. One milliliter of supernatant was subsequently centrifuged for 10 min at 13,800 × ɡ and filtered through a 0.2 μm filter (Whatman, Uppsala, Sweden). Filtrates were transferred to 2.0 mL GC vials (Agilent, Santa Clara, CA, USA). The concentration of VFAs was analyzed using a GC (6890N, Agilent, Santa Clara, CA, USA) equipped with a HP-INNOWax column (30 m × 0.25 mm × 0.25 μm; Agilent, Santa Clara, CA, USA) and a flame ionization detector (FID). The sample injection volume was 0.2 μL with a 10:1 split ratio. The oven temperature was initially temperature of 80°C for 2 min, increasing to 120°C at 20°C/min, then to 205°C at 10°C/min, and finally held at 205°C for 2 min. The injection and detection ports were maintained at 250°C.

Phenols and indoles.

Slurry samples were centrifuged at 3,134 × ɡ for 20 min at 20°C, and then 4 mL of supernatant was mixed with 4 mL of chloroform (Merck, Darmstadt, Germany) and 60 μL of 4M sodium hydroxide solution (Sigma-Aldrich, St. Louis, MO, USA) in a 20 mL glass vial. The mixture was centrifuged at 3,134 × ɡ for 20 min at 20°C, and the chloroform layer was transferred to a 2.0 mL GC vial (Agilent, Santa Clara, CA, USA). Phenols and indoles were analyzed using a GC (6890N, Agilent, Santa Clara, CA, USA) equipped with a DB-1 column (30 m × 0.25 mm × 0.25 μm, Agilent, Santa Clara, CA, USA) and a FID. The sample injection volume was 2.0 μL with a 5:1 split ratio. The oven temperature was initially 40°C for 5 min, increasing to 230°C at 10°C/min, which was then held at 230°C for 2 min. The injection and detection ports were maintained at 250°C.

Bacterial community analysis

PCR amplification for bar-coded pyrosequencing.

Total genomic DNA from slurry was extracted using a Fast-DNA Spin Kit (MP Bio, Santa Ana, CA, USA) according to the manufacturer’s instructions. Humic acid interferes with PCR amplification was removed using a Power-Clean DNA Clean-Up Kit (MP Bio, Santa Ana, CA, USA). Bacterial 16S ribosomal RNA (16S rRNA) genes around 500–700 bp long containing V1 to V3 of the variable region were amplified using primer set 27F (5'-adaptor 2-AC-GAG TTT GAT CMT GGC TCA G-3') and 518R (5'-adaptor 1-AC-X-WTT ACC GCG GCT GCT GG-3') where “X” denotes unique 7 to 11 bar-code sequences inserted between the 454 Life Sciences adaptor A sequence and the common linker, AC [21]. PCR amplification conditions were one cycle of 95°C for 5 min, followed by 30 cycles of 95°C for 30 sec, 55°C for 30 sec and 72°C for 30 sec, and finally one cycle of 72°C for 7 min.

Pyrosequencing and data analysis.

Pyrosequencing was performed by ChunLab (Seoul, Korea) using a 454 FLX Titanium System (Roche, Pleasanton, CA, USA). Sequencing reads were assigned to specific samples based on their unique barcodes. Then barcode, linker and PCR primer sequences at both ends were removed from the original sequencing reads. The final pyrosequencing reads for subsequent analysis were selected by a filtering process including only reads containing >300 base pairs and an average quality score >25. Taxonomic assignment of the bacterial high quality reads was performed using the EzTaxon-e database [22] and a robust global pairwise sequencing alignment, coupled with the BLAST search tool ( Sequences that could be matched to the EzTaxon-e database at the species level (>97%) were subjected to a secondary process to check for chimeric sequences using the UCHIME program [23]. Operational taxonomic units (OTUs) were generated using the CD-HIT program at a 97% similarity level. The Shannon-Weaver diversity index, Chao1 richness index and Goods library coverage were calculated using the Mothur package [24].

Statistical analysis

All experimental data including those concerning odorous compounds and bacterial communities were subjected to analysis of variance for a completely randomized design using the general linear model procedures of SAS software [25]. Significant differences among treatments were compared with Duncan’s multiple range tests [26]. Hierarchical clustering, principal component analysis (PCA) and principal coordinate analysis (PCoA) were performed to examine any grouping dependent upon the storage period. Pearson’s correlation coefficient was used to determine the link between bacterial genera and odorous compounds. Multiple statistical comparisons were evaluated using the data for odorous compounds and the bacterial community in the SAS program [25]. The threshold for significance was P<0.05 for all measured variables.


Concentration of odorous compounds in pig slurry by storage period

The effects of a storage period up to 6 weeks on phenols, indoles and VFAs concentration in pig slurry are shown in Table 1. The concentrations of odorous compounds were dramatically changed at 2 weeks-post storage period. Comparing 2 weeks storage to 4 weeks, phenols levels were decreased (P<0.05) by 18% (from 187.40 to 153.79 mg/L), and indoles levels were increased (P<0.05) by 44% (from 4.11 to 7.29 mg/L). SCFAs levels were decreased (P<0.05) by 18% (from 9,185 to 7,516 mg/L), and BCFAs levels were increased (P<0.05) by 24% (from 1,167 to 1,444 mg/L).

Table 1. Concentration of odorous compounds in pig slurry by storage period.

Changes in bacterial community composition in pig slurry by storage period

Pyrosequencing data.

Changes in the bacterial community structure during 6 weeks slurry storage period were analyzed by the multiplex bar-coded pyrosequencing technique based on 16S rRNA gene sequences (Table 2). After removal of low quality reads from the sequencing data, the total numbers of reads were 6,078 (0 weeks), 4,141 (2 weeks), 3,658 (4 weeks) and 4,000 (6 weeks), respectively. Although the number of bacterial reads was not significantly different after 2 weeks of storage, the number of OTUs, the Shannon-Weaver index and Chao 1 index were decreased (P<0.05) at 2 weeks-post storage.

Table 2. Pyrosequencing data in bacterial community composition in pig slurry by storage period.

Bacterial taxonomic composition.

Taxonomic pyrosequencing profiles of bacterial communities in pig slurry are shown in Figs 1 (phylum level) and 2 (genus level). The eight major phyla among a total of 27 included Firmicutes, Spirochaetes, Bacteroidetes, Tenericutes, Cloacamonas_p, Proteobacteria, Lentisphaerae and Actinobacteria (Fig 1). The relative abundance of Firmicutes and Lentisphaerae was reduced (P<0.05) after 2 weeks of storage. Bacteroidetes consistently decreased (P<0.05) during the 6 weeks storage period. However, Cloacamonas_p was drastically increased (P<0.05) after 2 weeks of storage.

Fig 1. Bacterial taxonomic composition of phylum level in pig slurry by storage period.

Sequences were classified using the EzTaxon-e database with an 80% confidence threshold. “wk” is an abbreviation to weeks.

Fig 2. Bacterial taxonomic composition of genus level in pig slurry by storage period.

Bacterial genus were classified at a cut off level of >0.5% relative abundance and grouped according to phylum level. “wk” is an abbreviation to weeks.

At the genus level (Fig 2), a total of 514 bacterial genera were represented in the sequences; they could be classified by phylum as: 253 Firmicutes, 70 Proteobacteria, 64 Bacteroidetes, 29 Actinobacteria, 23 Tenericutes, 21 Lentisphaerae, 7 Cloacamonas_p and 7 Spirochaetes. Altogether, there were 305 Gram-positive bacterial genera and 169 Gram-negative genera. Among the dominant genera, Clostridiales_uc_g, EU470107_g, Ruminococcaceae_uc, Lactobacillus, Turicibacter and HQ716403_g of the phylum Firmicutes, and Bacteroidales_uc_g, Bacteroides, Anaerocella_f_uc and Porphyromonadaceae_uc of the phylum Bacteroidetes were decreased (P<0.05) for 6 weeks of storage. Within these genera, Clostridiales_uc_g, Bacteroides and Porphyromonadaceae_uc were decreased (P<0.05) by 2 weeks. Sphaerochaeta of the phylum Spirochaetes, Cloacamonas_f_uc of Cloacaminas_p, GU454936_g of Firmicutes, AC160630_g of Bacteroidetes, Acholeplasmatales_uc_g and Mollicutes_uc_g of Tenericutes, and Advenella of Proteobacteria were increased (P<0.05) for 6 weeks of storage. Within these genera, Cloacamonas_f_uc, GU454936_g, AC160630_g, Acholeplasmatales_uc_g and Mollicutes_uc_g were drastically increased (P<0.05) after 2 weeks.

Statistical comparison of odorous compounds and bacterial community compositions in pig slurry by storage period

Changes in the bacterial compositions and concentrations of odorous compounds during the 6 weeks storage were graphically summarized using hierarchical clustering, PCA and PCoA in Figs 3 and 4. Two clusters were divided into one group of 0 and 2 weeks and another group of 4 and 6 weeks.

Fig 3. Hierarchical clustering result showing the group by storage period.

The clustering data was constructed using the concentrations of odorous compounds and the values of relative abundances in bacterial genus level. The scale bars represent the distance between clusters. “wk” is an abbreviation to weeks.

Fig 4. (A) Principal component analysis (PCA) and (B) principal coordinate analysis (PCoA) plot result showing the group by storage period.

These data were created using the concentrations of odorous compounds and the values of relative abundances in bacterial genus level. “wk” is an abbreviation to weeks.

Correlation between odorous compound production and dynamics of the bacterial community

The interrelationships of various bacterial genera associated with odorous compounds analyzed in the current study is shown in Table 3. Compared with other bacterial genera, Bacteroides, Porphyromonadaceae_uc_g, AC160630_g, Acholeplasmatales_uc_g, Mollicutes_uc_g and Cloacamonas_f_uc showed relatively greater (P<0.05) correlation coefficient values with odorous compounds. There was a positive correlation between phenol, p-cresol, acetic acid, propionic acid and butyric acid with Bacteroides and Porphyromonadaceae_uc_g, whereas there was a negative correlation with AC160630_g, Acholeplasmatales_uc_g, Mollicutes_uc_g and Cloacamonas_f_uc. Indole, skatole, iso-butyric acid and iso-valeric acid were shown to have the opposite correlations with these genera.

Table 3. Correlation coefficient (R) values for the relative abundance of bacterial genera with the odorous compounds in pig slurry.


The generation of odorous compounds from pig house is contributed to bacterial fermentation within the gastrointestinal tract of the pigs and the slurry in pit under the floor of the pig pen. Bacteria utilize undigested dietary materials, endogenous compounds and dead bacterial cells in slurry [17], and thereby generate various odorous compounds. Nitrogen is the major precursor of odorous compounds [28]. Gut bacteria obtain energy for maintaining homeostasis and growth from carbohydrates, and for building blocks from proteins included in the pig diet and endogenous materials. Availability of carbohydrate is very low while pig slurry is stored in the pit, thus protein becomes the main energy source for maintaining bacterial homeostasis [29]. In these circumstances, generation of offensive odorous compounds is augmented during bacteria-mediated amino acid metabolism [30]. This study was performed to identify the various odorous compounds generated from pig slurry stored for different periods, and their correlation with the bacterial taxonomic composition.

Phenols and indoles are produced during bacterial metabolism of tyrosine and tryptophan, respectively, in stored manure [27]. They are absorbed via epithelial cells of the large intestine, conjugated with glucuronic acid and detoxified to glucuronides in the liver. Glucuronides are excreted via urine and then hydrolyzed to phenols and indoles by fecal β–glucuronidase [31]. For this reason, combining the feces and urine results in raising the content of free phenolic or indolic compounds. They have a low odor detection threshold and relatively high in odor intensity and nuisance [28, 32]. The odor detection threshold is the lowest concentration of a certain odorous compound that is perceivable with the human sense of smell [33]. The detection thresholds for phenol, p-cresol, indole and skatole are 0.11, 0.00186, 0.0000316 and 0.000562 ppm, respectively [11, 3436]. Among the volatile organic compounds (VOCs) emitted from pig buildings, manure storage sites and land application, p-cresol, indole and skatole account for more than 90% of the odor activity value. In particular, p-cresol is most responsible for the overall odor impact from the VOCs emission [3739].

In this study, dramatic changes in the contents of phenols and indoles were detected at 2 weeks in stored slurry. Results from others have shown that a decrease in the concentration and emission rate of phenols started at about 36-day, but the indoles level was remained constant over 71-day storage period [40]. Ziemer, Kerr [41] reported that increased level of phenols and decreased level of indoles were detected after 8 weeks of storage, which correlated with a decrease in the pH. Phenols are formed at low pH, but indoles are accumulated at high pH [4244]. Therefore, decreased level of phenols and consistently high indoles level might relate to increased pH during slurry storage in our current study. In addition, phenol and p-cresol are degraded by aerobic bacteria [45], and phenol is also decomposed to CO2 by anaerobic bacteria in stored pig waste [46]. Bacterial decomposition is associated with bacterial growth when carbohydrate sources become limited as the storage period passes.

SCFAs are a product of carbohydrate fermentation, whereas BCFAs are a product of protein fermentation [27]. SCFAs are an important metabolic fuel for bacterial growth. SCFAs are absorbed into animal blood, transported to organs and tissues, and subsequently metabolized to energy sources for gut bacteria [27]. Decreased levels of SCFAs and increased BCFAs levels reflects a decrease in energy source availability in pig manure [40]. In the present study, SCFAs levels were decreased, but BCFAs levels increased in slurry stored for 6 weeks. VFAs concentrations dramatically changed after 2 weeks of storage. Miller and Varel [47] reported that a decrease in SCFAs (13%) and an increase in BCFAs (39%) occurred after 37 days of storage of pig slurry. Levels of VFAs are also affected by slurry pH; the BCFAs levels were increased by 67% at high pH (5.5 compared with 6.8) [48].

Analyzing the relationship between bacteria and their biotopes is an important step to understanding the mechanism of accumulation of odorous compounds produced by bacterial fermentation. The bacterial community in pig slurry was previously characterized by culture methods [4951]. Recently, multiplex bar-coded pyrosequencing techniques based on 16S rRNA genes have been used to identify members of the bacterial community comprising as little as 1% of the total population [7, 17, 52].

According to previous study [53], the bacterial community changed in pig slurry stored for 3 weeks; the number and diversity of bacteria decreased. In the present study (Fig 1) the relative abundance of Firmicutes, Bacteroidetes and Lentisphaerae decreased but Cloacamonas_p increased after 2 weeks storage of pig slurry. Firmicutes and Bacteroidetes constituted the majority of gut bacteria. Bacterial strains belonging to these two phyla are used as a phylogenetic markers because their relative abundance is easily influenced by fermentation conditions [54]. At the genus level, Clostridiales_uc_g, Bacteroides and Porphyromonadaceae_uc decreased but Cloacamonas_f_uc_g, GU454936_g, AC160630_g, Acholeplasmatales_uc_g and Mollicutes_uc_g increased after 2 weeks storage of pig slurry (Fig 2). Differences in the dominant bacterial groups might stem from the environmental changes in pig slurry during the storage period. The pH is usually higher in stored manure (pH 8~10) than fresh manure (pH 6~7) [17]. Some strains of Bacteroidetes are detected with low abundance at increased pH [55]. Strains of the phylum Cloacamonas_p increased during waste treatment utilizing livestock manure [56]. These bacteria that use amino acids as an energy source could be increased because of the low levels of available carbohydrate for bacterial growth in stored manure [29]. PCA and PCoA score plots (Fig 4) support this hypothesis, showing that bacterial groups and odorous compound levels were changed after 2 weeks of storage.

Bacterial composition in pig slurry is correlated with the content of odorous compounds produced. In the current study, phenol, p-cresol, acetic acid, propionic acid and butyric acid displayed a strong positive correlation with Bacteroides and Porphyromonadaceae_uc_g. Bacteroides have previously been isolated from manure stored in deep pits [57, 58]. These bacterial species are able to secrete various enzymes and decompose a wide range of carbohydrates and proteins [5961]. Among these enzymes, β-glucuronidase, which hydrolyses glucuronides to phenols and indoles, is produced at high levels [62]. In addition, hydrophobic molecules including phenols, indoles and their precursors could effectively transfer to the bacterial cell wall of Bacteroides [48, 63]. As a result, Bacteroides is related to changes in the levels of phenols and indoles in pig slurry. Most members of the family Porphyromonadaceae, including Porphyromonadaceae_uc_g, are saccharolytic excluding several proteolytic species [6466]. Major end products of these strains are acetic, butyric, propionic and lactic acid, with iso-butyric acid as a minor product [64, 6769]. Some strains are negatively related to indole formation [70].

Indole, skatole, iso-butyric acid and iso-valeric acid showed a positive correlation with AC160630_g, Acholeplasmatales_uc_g, Mollicutes_uc_g and Cloacamonas_f_uc_g. Acholeplasmatales_uc_g and Mollicutes_uc_g are members of the Mollicutes class in the phylum Tenericutes, Gram-positive and facultatively anaerobic [71]. The class Mollicutes arose from a Clostridium and descended from the Streptococcus/Lactobacillus group. Clostridium produces BCFAs and indoles from protein degradation [72, 73]. Lactobacillus strains convert indole-3-acetic acid, which is tryptophan metabolite formed by Clostridium, to skatole [42]. The Mollicutes class also produces some organic acids in anaerobic conditions. The members of the order Acholeplasmatales are the most abundant among the Mollicutes class and ferment glucose to produce acetic and lactic acid [74]. In addition, many strains of the Acholeplasmatales order are capable of synthesizing other fatty acids by utilizing acetate and butyrate [75]. Mollicutes strains are positively associated with the synthesis of valerate, iso-butyrate and iso-valerate, and can use amino acids as a carbon source [76]. The phylum Cloacamonas_p, including the genus Cloacamonas_f_uc_g, is a Candidatus taxon belonging to a subdominant bacterial group called WWE1 (Waste Water of Evry 1) and discovered by molecular inventories of an anaerobic digester [77]. WWE1 members could hydrolyze the cellulose via the release of extracellular cellulase. Cloacamonas spp. is syntrophic amino acid metabolizer, which could ferment a highly enriched hydrolysis product resulting from the hydrolytic activity of other bacteria [78] and produces fatty acids by phenol degradation [79]. Therefore, these bacterial strains are capable of the degradation of proteins for growth in the hypo-carbohydrate condition. In the present study, increased abundance of members of the phylum Tenericutes and Cloacamonas in stored slurry can be explain the protein availability for growth in the hypo-carbohydrate condition and then the high indoles and BCFA levels, which would be derived mainly from amino acid fermentation.


Our results demonstrate that the storage period of pig slurry in a pit significantly affects the composition of odorous compounds produced as well as the bacterial community. Levels of odorous compounds were dramatically changed after 2 weeks of slurry storage. Phenols and SCFAs decreased, whereas indoles and BCFAs increased in the pig slurry. Accumulation of indoles and BCFAs is associated with increased pH during slurry storage and showed a strong positive correlation with members of the taxa Tenericutes and Cloacamonas. Tenericutes and Cloacamonas strains use proteins as a carbon source for growth and predominate in various environmental conditions. Our current study has significant value in identifying the causes of odor from pig houses. Based on our results, it is desirable that pig slurry needs to be discharged every 2 weeks to reduce the odor in pig house. Further investigation is necessary to control bacterial growth and identify fermentation patterns to increase the efficiency of odor reduction.


This study was carried out with the support of “Practical use of odor reducing agent developed to symbiotic technology for pigpen (Project title: Practical research of odor reducing agent to add in pigpen slurry, Project No. PJ01026302)” by National Institute of Animal Science, Rural Development Administration, Republic of Korea.

Author Contributions

  1. Conceptualization: OHH SBC SKP.
  2. Data curation: OHH.
  3. Formal analysis: OHH SBC SKP.
  4. Funding acquisition: OHH.
  5. Investigation: OHH SBC SKP.
  6. Methodology: OHH.
  7. Project administration: OHH SBC.
  8. Resources: OHH SBC SKP DWH SRL JHK.
  9. Supervision: SBC SKP.
  10. Validation: OHH SBC SKP.
  11. Visualization: OHH.
  12. Writing – original draft: OHH.
  13. Writing – review & editing: OHH SKP.


  1. 1. MAFRA. Ministry of agriculture, food and rural affairs annual report 2013. Republic of Korea: Ministry of Agriculture Food and Rural Affairs, 2013.
  2. 2. MAFRA. Ministry of agriculture, food and rural affairs annual report 2010. Republic of Korea: Ministry of Agriculture Food and Rural Affairs, 2010.
  3. 3. MEV. Law on offensive odor control. Republic of Korea: Ministry of Environment, 2012.
  4. 4. MEV. Law on livestock excretions management and usage. Republic of Korea: Ministry of Environment, 2011.
  5. 5. Zhu J. A review of microbiology in swine manure odor control. Agr Ecosyst Environ. 2000; 78(2): 93–106.
  6. 6. Williams A, Evans M. Storage of piggery slurry. Agr Wastes. 1981; 3(4): 311–321.
  7. 7. Snell-Castro R, Godon J-J, Delgenès J-P, Dabert P. Characterisation of the microbial diversity in a pig manure storage pit using small subunit rDNA sequence analysis. FEMS Microbiol Ecol. 2005; 52(2): 229–242. pmid:16329909
  8. 8. Kai P, Schäfer A. Identification of key odour components in pig house air using hyphenated gas chromatography olfactometry. Agric Eng Int: GIGR Journal. 2004; VI.
  9. 9. Trabue S, Kerr B, Bearson B, Ziemer C. Swine odor analyzed by odor panels and chemical techniques. J Environ Qual. 2011; 40(5): 1510–1520. pmid:21869513
  10. 10. Trabue S, Scoggin K, McConnell L, Maghirang R, Razote E, Hatfield J. Identifying and tracking key odorants from cattle feedlots. Atmos Environ. 2011; 45(25): 4243–4251.
  11. 11. O'neill D, Phillips V. A review of the control of odour nuisance from livestock buildings: Part 3, properties of the odorous substances which have been identified in livestock wastes or in the air around them. J Agr Eng Res. 1992; 53: 23–50.
  12. 12. Mackie R. Microbial production of odor components. Proceedings of the International Round table on Swine Odor Control. 1994: 13–15.
  13. 13. Zahn J, Hatfield J, Do Y, DiSpirito A, Laird D, Pfeiffer R. Characterization of volatile organic emissions and wastes from a swine production facility. J Environ Qual. 1997; 26(6): 1687–1696.
  14. 14. Rappert S, Müller R. Odor compounds in waste gas emissions from agricultural operations and food industries. Waste Manage. 2005; 25(9): 887–907.
  15. 15. Cummings J, Macfarlane G. The control and consequences of bacterial fermentation in the human colon. J Appl Bacteriol. 1991; 70(6): 443–459. pmid:1938669
  16. 16. Klose V, Bayer K, Bruckbeck R, Schatzmayr G, Loibner A-P. In vitro antagonistic activities of animal intestinal strains against swine-associated pathogens. Vet Microbiol. 2010; 144(3): 515–521.
  17. 17. Whitehead TR, Cotta MA. Characterisation and comparison of microbial populations in swine faeces and manure storage pits by 16S rDNA gene sequence analyses. Anaerobe. 2001; 7(4): 181–187.
  18. 18. Leung K, Topp E. Bacterial community dynamics in liquid swine manure during storage: molecular analysis using DGGE/PCR of 16S rDNA. FEMS Microbiol Ecol. 2001; 38(2–3): 169–177.
  19. 19. NIAS. Korean feeding standard, swine. Republic of Korea: National institute of animal science 2012.
  20. 20. Zhu J, Bundy DS, Li XW, Rashid N. Reduction of odor and volatile substances in pig slurries by using pit additives. Journal of Environmental Science and Health Part A-Environmental Science and Engineering & Toxic and Hazardous Substance Control. 1996; 31(10): 2487–2501. WOS:A1996VW41600005.
  21. 21. Chun J, Kim KY, Lee J-H, Choi Y. The analysis of oral microbial communities of wild-type and toll-like receptor 2-deficient mice using a 454 GS FLX Titanium pyrosequencer. BMC Microbiol. 2010; 10(1): 101.
  22. 22. Kim O-S, Cho Y-J, Lee K, Yoon S-H, Kim M, Na H, et al. Introducing EzTaxon-e: a prokaryotic 16S rRNA gene sequence database with phylotypes that represent uncultured species. Int J Syst Evol Micr. 2012; 62(Pt 3): 716–721.
  23. 23. Edgar RC, Haas BJ, Clemente JC, Quince C, Knight R. UCHIME improves sensitivity and speed of chimera detection. Bioinformatics. 2011; 27(16): 2194–2200. pmid:21700674
  24. 24. Schloss PD, Westcott SL, Ryabin T, Hall JR, Hartmann M, Hollister EB, et al. Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Appl Environ Microb. 2009; 75(23): 7537–7541.
  25. 25. SAS. SAS/STAT software, version 9.1. SAS Institute Inc., Cary, N.C. USA.; 2002.
  26. 26. Duncan DB. Multiple range and multiple F tests. Biometrics. 1955; 11(1): 1–42.
  27. 27. Mackie RI, Stroot PG, Varel VH. Biochemical identification and biological origin of key odor components in livestock waste. J Anim Sci. 1998; 76(5): 1331–1342. pmid:9621939.
  28. 28. Le PD, Aarnink AJ, Ogink NW, Becker PM, Verstegen MW. Odour from animal production facilities: its relationship to diet. Nutr Res Rev. 2005; 18(1): 3–30. pmid:19079892.
  29. 29. Le P, Aarnink A, Jongbloed A, Van der Peet-Schwering C, Ogink N, Verstegen M. Interactive effects of dietary crude protein and fermentable carbohydrate levels on odour from pig manure. Livest Sci. 2008; 114(1): 48–61.
  30. 30. Willig S, Lösel D, Claus R. Effects of resistant potato starch on odor emission from feces in swine production units. J Agr Food Chem. 2005; 53(4): 1173–1178.
  31. 31. Roustan J, Aumaitre A, Bernard C. Tentative measurements of animal manure odor. London: Applied Science Publishers; 1980.
  32. 32. Jensen BB, Jørgensen H. Effect of dietary fiber on microbial activity and microbial gas production in various regions of the gastrointestinal tract of pigs. Appl Environ Microb. 1994; 60(6): 1897–1904.
  33. 33. Zhang S, Cai L, Koziel JA, Hoff SJ, Heathcote KY, Jacobson L, et al., editors. Odor and Odorous Chemical Emissions from Animal Buildings: Part 5–Correlations between Odor Intensities and Chemical Concentrations (gc-ms/o). International Symposium on Air Quality and Manure Management for Agriculture Conference Proceedings, 13–16 September 2010, Dallas, Texas; 2010: American Society of Agricultural and Biological Engineers.
  34. 34. Sutton AL, Kephart KB, Verstegen MW, Canh TT, Hobbs PJ. Potential for reduction of odorous compounds in swine manure through diet modification. J Anim Sci. 1999; 77(2): 430–439. pmid:10100673
  35. 35. Hammond EG, Heppner C, Smith R. Odors of swine waste lagoons. Agr Ecosyst Environ. 1989; 25(2): 103–110.
  36. 36. Schiffman SS, Bennett JL, Raymer JH. Quantification of odors and odorants from swine operations in North Carolina. Agr Forest Meteorol. 2001; 108(3): 213–240.
  37. 37. Parker DB, Gilley J, Woodbury B, Kim K-H, Galvin G, Bartelt-Hunt SL, et al. Odorous VOC emission following land application of swine manure slurry. Atmos Environ. 2013; 66: 91–100.
  38. 38. Trabue SL, Scoggin KD, Li H, Burns R, Xin H. Field sampling method for quantifying odorants in humid environments. Envir Sci Tech. 2008b; 42(10): 3745–3750.
  39. 39. Trabue SL, Scoggin K, Mitloehner F, Li H, Burns R, Xin H. Field sampling method for quantifying volatile sulfur compounds from animal feeding operations. Atmos Environ. 2008a; 42(14): 3332–3341.
  40. 40. Hobbs P, Misselbrook T, Cumby T. Production and emission of odours and gases from ageing pig waste. J Agr Eng Res. 1999; 72(3): 291–298.
  41. 41. Ziemer CJ, Kerr BJ, Trabue SL, Stein H, Stahl DA, Davidson SK. Dietary protein and cellulose effects on chemical and microbial characteristics of swine feces and stored manure. J Environ Qual. 2009; 38(5): 2138–2146. pmid:19704156
  42. 42. Jensen MT, Cox RP, Jensen BB. 3-Methylindole (skatole) and indole production by mixed populations of pig fecal bacteria. Appl Environ Microbiol. 1995; 61(8): 3180–3184. pmid:7487051; PubMed Central PMCID: PMC167595.
  43. 43. Smith MB, March J. March's advanced organic chemistry: reactions, mechanisms, and structure: John Wiley & Sons; 2007.
  44. 44. Wu JJ, Park S-h, Hengemuehle SM, Yokoyama MT, Person HL, Gerrish JB, et al. The use of ozone to reduce the concentration of malodorous metabolites in swine manure slurry. J Agr Eng Res. 1999; 72(4): 317–327.
  45. 45. Ishaque M, Bisaillon J-G, Beaudet R, Sylvestre M. Degradation of phenolic compounds by microorganisms indigenous to swine waste. Agr Wastes. 1985; 13(3): 229–235.
  46. 46. Boopathy R. Isolation and characterization of a phenol-degrading, sulfate-reducing bacterium from swine manure. Bioresource Technol. 1995; 54(1): 29–33.
  47. 47. Miller D, Varel V. Swine manure composition affects the biochemical origins, composition, and accumulation of odorous compounds. J Anim Sci. 2003; 81(9): 2131–2138. pmid:12968686
  48. 48. Smith EA, Macfarlane GT. Enumeration of amino acid fermenting bacteria in the human large intestine: effects of pH and starch on peptide metabolism and dissimilation of amino acids. FEMS Microbiol Ecol. 1998; 25(4): 355–368.
  49. 49. Hobson P, Shaw B. The bacterial population of piggery-waste anaerobic digesters. Water Res. 1974; 8(8): 507–516.
  50. 50. Iannotti E, Fischer J, Sievers D. Characterization of bacteria from a swine manure digester. Appl Environ Microb. 1982; 43(1): 136–143.
  51. 51. Spoelstra SF. Microbial aspects of the formation of malodorous compounds in anaerobically stored piggery wastes: Landbouwhogeschool te wageningen.; 1978.
  52. 52. Hamady M, Walker JJ, Harris JK, Gold NJ, Knight R. Error-correcting barcoded primers allow hundreds of samples to be pyrosequenced in multiplex. Nat Methods. 2008; 5(3): 235.
  53. 53. Peu P, Brugère H, Pourcher A-M, Kérourédan M, Godon J-J, Delgenès J-P, et al. Dynamics of a pig slurry microbial community during anaerobic storage and management. Appl Environ Microb. 2006; 72(5): 3578–3585.
  54. 54. Ramakrishna BS. Role of the gut microbiota in human nutrition and metabolism. J Gastroen Hepatol. 2013; 28(S4): 9–17.
  55. 55. Goux X, Calusinska M, Lemaigre S, Marynowska M, Klocke M, Udelhoven T, et al. Microbial community dynamics in replicate anaerobic digesters exposed sequentially to increasing organic loading rate, acidosis, and process recovery. Biotechnol Biofuels. 2015; 8(1): 1.
  56. 56. Solli L, Håvelsrud OE, Horn SJ, Rike AG. A metagenomic study of the microbial communities in four parallel biogas reactors. Biotechnol Biofuels. 2014; 7(1): 1–15.
  57. 57. Cotta MA, Whitehead TR, Zeltwanger RL. Isolation, characterization and comparison of bacteria from swine faeces and manure storage pits. Environ Microbiol. 2003; 5(9): 737–745. pmid:12919409
  58. 58. Spoelstra SF. Enumeration and isolation of anaerobic microbiota of piggery wastes. Appl Environ Microb. 1978; 35(5): 841–846.
  59. 59. Gaskins HR. Intestinal bacteria and their influence on swine growth. Washington, DC: CRC Press 2001. 609–630 p.
  60. 60. Riepe SP, Goldstein J, Alpers DH. Effect of secreted Bacteroides proteases on human intestinal brush border hydrolases. J Clin Invest. 1980; 66(2): 314. pmid:6995483
  61. 61. Salyers AA. Energy sources of major intestinal fermentative anaerobes. Am J Clin Nutr. 1979; 32(1): 158–163. pmid:367143
  62. 62. McBain A, Macfarlane G. Ecological and physiological studies on large intestinal bacteria in relation to production of hydrolytic and reductive enzymes involved in formation of genotoxic metabolites. J Med Microbiol. 1998; 47(5): 407–416. pmid:9879941
  63. 63. Van der Meulen R, Camu N, Van Vooren T, Heymans C, De Vuyst L. In vitro kinetic analysis of carbohydrate and aromatic amino acid metabolism of different members of the human colon. Int J Food Microbiol. 2008; 124(1): 27–33. pmid:18396344
  64. 64. Sakamoto M, Takagaki A, Matsumoto K, Kato Y, Goto K, Benno Y. Butyricimonas synergistica gen. nov., sp. nov. and Butyricimonas virosa sp. nov., butyric acid-producing bacteria in the family ‘Porphyromonadaceae’isolated from rat faeces. Int J Syst Evol Micr. 2009; 59(7): 1748–1753.
  65. 65. Ziganshin AM, Schmidt T, Scholwin F, Il’inskaya ON, Harms H, Kleinsteuber S. Bacteria and archaea involved in anaerobic digestion of distillers grains with solubles. Appl Microbiol Biot. 2011; 89(6): 2039–2052.
  66. 66. Qiu Y-L, Kuang X-Z, Shi X-S, Yuan X-Z, Guo R-B. Paludibacter jiangxiensis sp. nov., a strictly anaerobic, propionate-producing bacterium isolated from rice paddy field. Arch Microbiol. 2014; 196(3): 149–155. pmid:24419224
  67. 67. Ueki A, Akasaka H, Suzuki D, Ueki K. Paludibacter propionicigenes gen. nov., sp. nov., a novel strictly anaerobic, Gram-negative, propionate-producing bacterium isolated from plant residue in irrigated rice-field soil in Japan. Int J Syst Evol Micr. 2006; 56(1): 39–44.
  68. 68. Hofstad T, Olsen I, Eribe ER, Falsen E, Collins MD, Lawson PA. Dysgonomonas gen. nov. to accommodate Dysgonomonas gadei sp. nov., an organism isolated from a human gall bladder, and Dysgonomonas capnocytophagoides (formerly CDC group DF-3). Int J Syst Evol Micr. 2000; 50(6): 2189–2195.
  69. 69. Song Y, Liu C, Lee J, Bolaňos M, Vaisanen M-L, Finegold SM. “Bacteroides goldsteinii sp. nov.” Isolated from Clinical Specimens of Human Intestinal Origin. J Clin Microbiol. 2005; 43(9): 4522–4527. pmid:16145101
  70. 70. Andrea IS, Sanz JL, Stams A. Microbacter margulisiae gen. nov., sp. nov., a novel propionigenic bacterium isolated from sediments of an acid rock drainage pond. Int J Syst Evol Micr. 2014; 64: 3936–3942.
  71. 71. Gundersen DE, Lee I-M, Rehner SA, Davis RE, Kingsbury DT. Phylogeny of mycoplasmalike organisms (phytoplasmas): a basis for their classification. J Bacteriol. 1994; 176(17): 5244–5254. pmid:8071198
  72. 72. Elsden SR, Hilton MG. Volatile acid production from threonine, valine, leucine and isoleucine by clostridia. Arch Microbiol. 1978; 117(2): 165–172. pmid:678022
  73. 73. Chung KT, Anderson GM, Fulk GE. Formation of indoleacetic acid by intestinal anaerobes. J Bacteriol. 1975; 124(1): 573–575. pmid:1236846
  74. 74. Lazarev V, Levitskii S, Basovskii YI, Chukin M, Akopian T, Vereshchagin V, et al. Complete genome and proteome of Acholeplasma laidlawii. J Bacteriol. 2011; 193(18): 4943–4953. pmid:21784942
  75. 75. Kube M, Siewert C, Migdoll AM, Duduk B, Holz S, Rabus R, et al. Analysis of the complete genomes of Acholeplasma brassicae, A. palmae and A. laidlawii and their comparison to the obligate parasites from'Candidatus Phytoplasma'. J Mol Microb Biotech. 2013; 24(1): 19–36.
  76. 76. Mao S, Zhang R, Wang D, Zhu W. The diversity of the fecal bacterial community and its relationship with the concentration of volatile fatty acids in the feces during subacute rumen acidosis in dairy cows. BMC Vet Res. 2012; 8(1): 237.
  77. 77. Pelletier E, Kreimeyer A, Bocs S, Rouy Z, Gyapay G, Chouari R, et al. “Candidatus Cloacamonas acidaminovorans”: genome sequence reconstruction provides a first glimpse of a new bacterial division. J Bacteriol. 2008; 190(7): 2572–2579.
  78. 78. Limam RD, Chouari R, Mazeas L, Wu TD, Li T, Grossin‐Debattista J, et al. Members of the uncultured bacterial candidate division WWE1 are implicated in anaerobic digestion of cellulose. MicrobiologyOpen. 2014; 3(2): 157–167. pmid:24497501
  79. 79. Ju F, Zhang T. Novel microbial populations in ambient and mesophilic biogas-producing and phenol-degrading consortia unraveled by high-throughput sequencing. Microb Ecol. 2014; 68(2): 235–246. pmid:24633337