Bioenergy from anaerobic degradation of lipids in palm oil mill effluent

  • Anwar Ahmad
  • Rumana Ghufran
  • Zularisam Abd. Wahid
Open Access


Fossil fuels are the lifeblood of our society and for many others around the world. The environmental pollution due to the use of fossil fuels as well as their gradual depletion make it necessary to find alternative energy and chemical sources that are environmentally friendly and renewable. Palm oil mill effluent (POME), a strong wastewater from palm oil mills, has been identified as a potential source to generate renewable bioenergies through anaerobic digestion. Thus, it has received considerable attention as feedstock for producing various value added products such as methane gas, bio-plastic, organic acids, bio-compost, activated carbon, and animal feedstock. Lipids are one of the major organic pollutants in POME. Furthermore, waste lipids are ideal potential substrates for biogas production, since theoretically more methane can be produced, when compared with proteins or carbohydrates. The objective of this review paper is to disscuss the microbial communities involved in the anaerobic degradation of long chain fatty acid and bioenergies and by-products from POME. With these options (Renewable and sustainable bioenergies) we can help phase out our dependency on fossil fuels and find clean, efficient, sources of power.


Palm oil mill effluent (POME) Long chain fatty acids (LCFAs) Green house gas (GHG) β-oxidation Lipids 



Palm oil mill effluent


Long chain fatty acids


Greenhouse gases


Chemical oxygen demand


Biochemical oxygen demand


Carbon dioxide






Standard temperature and pressure




Continuous stirred tank reactor


Anaerobic baffled reactor


Up flow anaerobic sludge blanket


Expanded granular sludge blanket


Denaturating gradient gel electrophoresis


Up-flow anaerobic sludge fixed-film reactor


Modified anaerobic baffled reactor


Fluidized bed reactor




Poly hydroxy alkanoate


Poly-β-hydroxy butyric acid


Organic loading rate


Hydraulic retention time


Volatile fatty acid

1 Introduction

Over the last 30 years, Malaysian palm oil industry has grown rapidly and at present it is one of the largest agro-based industries in the world (Wong et al. 2002; Wu et al. 2010). Palm oil mill effluent is considered as one of the most polluting agro-industrial effluent due to its high values of COD and BOD. Today, the percolation of palm oil mill effluents into the waterways and ecosystems, remain a fastidious concern towards the public health and food chain interference (Foo and Hameed 2010). This can cause considerable environmental problems if discharged without effective treatment by polluting land, water and destroying aquatic biota (Cheng et al. 2010; Singh et al. 2011). Therefore, palm oil mills are required to treat their POME prior to discharging it into streams and rivers. In the process of palm oil milling, POME is mainly generated from sterilization and clarification of palm oil in which a large amount of steam and hot water are used (Zinatizadeh et al. 2006; Rupani et al. 2010). POME is acidic (pH 4–5), has discharge temperature of 80–90°C/50–60°C and is non toxic (since no chemicals are added during extraction), (Ahmad et al. 2003). The characteristics and the parameter limits for POME discharge into watercourses in Malaysia are summarized in Table 1. It is rich in organic matter such as proteins, carbohydrates and lipids along with nitrogenous compounds and minerals (Agamuthu and Tan 1985; Habib et al. 1997; Wu et al. 2007). Therefore, it can be reused for biotechnological means. The various effluent treatment schemes, which are currently used by the Malaysian palm oil industry, are listed in descending order: (a) anaerobic/facultative ponds (Rahim and Raj 1982; Wong 1980; Chan and Chooi 1982), (b) tank digestion and mechanical aeration, (c) tank digestion and facultative ponds, (d) decanter and facultative ponds, (e) physico-chemical and biological treatment (Andreasen 1982), and (f) evaporation (Ma 1999a, b) and a clarification coupled with filtration and aeration (UNEP 1994).
Table 1

Characteristics and parameter limits for POME discharge into water courses in Malaysia

Characteristics of POME

Parameter limits for watercourse discharge for POME

Major constituents of POME

Quantity (g/g dry sample)




Limits of dischargea







Crude protein





Crude lipid


Biological Oxygen Demand 3-days 30°Cb






Chemical Oxygen Demand





Total solids



Nitrogen-free extract


Suspended Solids




Total carotene


Volatile Solids




Oil and Grease





Ammoniacal Nitrogen





Total Nitrogen





Source: (Ma 1999a, b, 2000)

Source: EQA 1974 (Act 127) and Subsidiary Legislation 2002

Source: Habib et al. (1997)

aUnits in mg/l except pH and Temperature (°C)

bThe sample for BOD analysis is incubated at 30°C for 3 days

cValue of filtered sample

In an anaerobic process, the POME is degraded into methane, carbon dioxide and water, and the sequence of reactions included; hydrolysis, acidogenesis (including acetogenesis) and methanogenesis (Bitton 2005; Nwuche and Ugoji 2008; Nwuche and Ugoji 2010). In the process of hydrolysis the complex molecules (i.e. carbohydrates, lipids and proteins) are hydrolyzed into sugars, amino acids and fatty acids etc. by extra cellular enzymes of fermentative bacteria. In acidogenesis, these sugars, fatty acids and amino acids are converted into organic acids by means of acidogenic bacteria. These are further converted to acetate together with CO2 and hydrogen by acetogens. Finally the hydrogen is utilized by hydrogenotrophic methanogens while acetic acid and CO2 are utilized by acetoclastic methanogens to methane as a final product. Thus, anaerobic decomposition of organic matter involves the concerted action of several different metabolic groups of microorganisms (Demirel and Scherer 2008; Weiland 2010) to produce biogas that can be used to generate electricity and save fossil energy (Wong et al. 2009).

The compositions and concentrations of proteins, nitrogenous compounds, lipids and minerals found to be present in POME (Habib et al. 1997) are summarized in Table 2. Lipids are one of the major organic pollutants in POME. These compounds are glycerol, bonded to LCFAs, alcohols, and other groups by an ester or ether linkage. During anaerobic treatment, lipids are hydrolyzed into glycerol plus LCFAs by hydrolytic extracellular lipases. Glycerol is further degraded via acidogenesis while LCFAS are degraded through β-oxidation process (syntrophic acetogenesis) to acetate, H2 and CO2 (Jeris and McCarty 1965; Weng and Jeris 1976) and finally converted to CH4 or CO2 by methanogenesis (Komatsu et al. 1991; Stryer 1995). Lipid hydrolysis is generally faster than protein or carbohydrate hydrolysis and considered to be a rapid process in anaerobic digestion while subsequent β-oxidation proceeds rather slowly (Pavlostathis and Geraldogomez 1991; Hanaki et al. 1981; Angelidaki and Ahring 1995). After hydrolysis, LCFAS glycerol undergoes fermentation or acidogenesis. Since LCFA requires an external electron acceptor for oxidation, therefore, its degradation occurs during acetogenesis while mainly saturation or hydrogenation of unsaturated LCFAs takes place in this process. Mainly glycerol is also degraded to acetate, lactate and 1,3-propanediol (Batstone 2000). Proton-reducing acetogens degrade LCFAs in syntrophic association with hydrogen-utilizing methanogens and acetoclastic methanogens (Schink 1997).
Table 2

Fatty acids, amino acids and mineral contents of raw POME (Source: Habib et al. 1997)

Fatty acids (g/100 g lipid)

Quantity (g)

Chemical formula of fatty acids

Amino acids (g/100 g protein)

Chemical formula

Quantity (g)


Quantity (mm/g dry sample)

Capric acid (10:0)



Aspartic acid





Lauric acid (12:0)



Glutamic acid





Myristic acid (14:0)








Palmitic acid (16:0)








Heptadecanoic acid (17:0)








10-heptadecanoic acid (17:01)








Stearic acid (18:0)








Oleic acid (18:1n-9)








Lenoleic acid (18:2n-6)








Linolenic acid (18:3n-3)








Arachidic acid (20:0)








Eicosatrienoic acid (20:3n-6)








Arachidonic acid (20:4n-6)








Eicosapentaeonic acid (20:5n-3)








Behenic acid (22:0)












































Syntrophism is a special case of mutualistic interrelationship between two different microorganisms which together degrade some substances (and conserve energy doing it) which neither could do separately. Thus, this term was coined to describe the close cooperation of fatty acid-oxidizing, fermenting bacteria with hydrogen oxidizing methanogens (McInery et al. 1979) or of phototrophic green sulphur bacteria with chemotropic sulphur-reducing bacteria (Biebl and Pfening 1978). In most cases of syntrophism the nature of a syntrophic reaction involves H2 gas being produced by one partner and being consumed by the other.

Oxidation of butyric acid to acetic acid and H2 by the fatty acid-oxidizing Syntrophomonas is a good example of mutual dependence. Syntrophomonas does not grow in a pure culture on butyric acid as the energy released during butyric acid oxidation to acetic acid is highly unfavorable to the bacterium. But, if the hydrogen produced in the reaction is immediately utilized by a syntrophic partner (e.g. methanogen), Syntrophomonas grows luxuriantly in mixed-culture with the H2 consumer.
$$ {\text{Butyrate}}^{ - } + 2 {\text{H}}_{ 2} {\text{O}} \to 2 {\text{Acetate}}^{ - } + {\text{H}}^{ + } + 2 {\text{H}}_{ 2} + {\text{Energy }}\left( { + 4 8. 2 {\text{ Kj}}} \right) $$
Wastes or wastewaters with a high lipid-content present an attractive source for methane production because theoretically it has been found that 1.01 L of methane at STP can be produced from 1 g of oleate (unsaturated LCFA, C18:1), whereas only 0.37 L can be produced from 1 g of glucose (Kim et al. 2004). Thus, degradation of lipids produces more biogas with higher methane content as compared to proteins or carbohydrates (Table 3).
Table 3

Potential biogas production from different classes of substrates


Methanogenic reaction

Biogas (lg−1)

CH4 (%)


C50H90O6 + 24.5 H2O → 34.75 CH4 + 15.25 CO2




C6H10O5 + H2O → 3CH4 + 3CO2




C16H24O5N4 + 14.5 H2O → 8.25 CH4 + 3.75 CO2 + 4NH4 + + 4HCO3



Source: Alves et al. (2009)

This paper will provide a brief overview of the biochemistry, microbiology of anaerobic breakdown of polymeric materials especially of lipids to methane and the roles of the various microorganisms involved along with a brief description of bioenergies and biochemical from POME.

2 Food web of lipid anaerobic process

Anaerobic bioconversion of complex organic matter to methane consists of a series of microbiological processes. As shown in the Fig. 1, first step involves the hydrolysis of complex organic polymers (proteins, carbohydrates and lipids) into monomers by extracellular enzymes that are excreted by fermentative bacteria. Proteins are degraded to amino acids, carbohydrates to soluble sugars and lipids to glycerol and LCFAs by the concerted action of proteases, cellulases and lipases respectively. Some researchers suggested that lipid hydrolysis is generally faster than protein or carbohydrate hydrolysis, and considered to be a rapid process in anaerobic digestion (Pavlostathis and Geraldogomez 1991; Hanaki et al. 1981; Angelidaki and Ahring 1995), while some other researchers reported lipid hydrolysis as a rate limiting step (Beccari et al. 1996). According to some other researchers lipid hydrolysis was not usually the rate limiting step, the overall conversion rate was limited either by degradation of LCFAs or by the physical processes of dissolution and mass transfer of these acids (Rinzema 1993; Hanaki et al. 1987). The apparent discrepancy between the results reported by different researchers may be attributed to differences in the feed concentration and physico-chemical conditions in their experiments.
Fig. 1

Food web of methanogenic anaerobic digestion

After hydrolysis, LCFAs and glycerol undergo fermentation or acidogenesis. Although hydrolysis of lipids to glycerol and LCFAs occurs rapidly, subsequent LCFAs degradation via β-oxidation proceeds rather slowly (Pavlostathis and Geraldogomez 1991). Since LCFAs oxidation requires an external electron acceptor, therefore, undergo degradation during acetogenesis while unsaturated LCFAs get saturated in this process and glycerol degraded to acetate, lactate and 1,3 propandiol (Batstone 2000). Weng and Jeris (1976) suggested that degradation of unsaturated LCFAs starts with chain saturation, whereas other authors provide evidence that the direct β-oxidation of unsaturated LCFAs is feasible (Lalman and Bagley 2001; Lalman and Bagley 2000; Roy et al. 1986). However, it is still unknown whether saturated and unsaturated LCFAs are degraded by the same bacteria in methanogenic environments.

During syntrophic acetogenesis the products of acidogenesis are converted into methanogenic substrates i.e. acetate, hydrogen and carbondioxide for methanogenesis. The formation of methanogenic subsrate with H2 production is thermodynamically unfavourable unless the partial pressure of H2 in the medium is kept low by H2 consuming bacteria such as methanogens (McInerney et al. 1981). Thus, LCFA oxidation to acetate is possible only when the hydrogen partial pressure in the medium is kept low and, therefore, cooperation with hydrogen-consuming microorganisms is necessary (Table 4). For this oxidation, the partial pressure of hydrogen has to be decreased substantially to lower values (<10 Pa) than the ethanol because degradation of fatty acids to CH3COO and H2, or in the case of propionate, to CH3COO, H2 or CO2 is far more endergonic under standard conditions than ethanol oxidation (Table 5). Because of the diverse number of organisms involved in these reactions and their ability to perform other types of metabolisms such as fermentation or sulfate reduction (Schink 1997; McInerney 1992), the organisms that participate at this second step will be called syntrophic metabolizers. Table 5 summarizes some reactions involved in syntrophic metabolism. Normally all LCFAs and ethanol require syntrophic acetogenesis along with two pathways: (1) β-oxidation pathway (as the main pathway) and (2) \( \tilde{\omega } \)-oxidation pathway by carboxylation of far methyl carbon.
Table 4

Gibbs free energy changes at 25°C for the (possible) reactions involved in syntrophic conversion of oleate and palmitate during methanogenic decomposition (source: Sousa et al. 2007a, b, c)

Reaction No.



\( \Updelta {\text{G}}^{{ 0^{\prime } }} \) (KJ reaction−1)b

\( \Updelta {\text{G}}^{{ 0^{\prime } }} \) (KJ reaction−1)c


Oleate degradation

C18H33O 2 2−  + 16H2O → 9C2H3O2− + 15H2 + 8H+




Palmitate degradation

C16H31O2 2− + 14H2O → 8C2H3O2 2− + 14H2 +7H+




Hydrogenotrophic methanogens

4H2 + HCO3  + H+ → CH4 + 3H2O




Acetoclastic methanogens

C2H3O2  + H2O → HCO3  + CH4



Data were obtained or calculated from data reported by Thauer et al. (1977) and Lalman (2000)

aGibbs free energies (at 25°C) calculated under standard conditions (solute concentrations of 1 M and gas partial pressure of 105 Pa)

bGibbs free energies (at 25°C) for oleate/palmitate concentration of 1 mM, acetate concentration of 10 mM, and H2 partial pressure of 1 Pa

Table 5

Reactions involved in syntrophic metabolism

Reaction input

Reaction output

\( \Updelta {\text{G}}^{{ 0^{\prime } }} \) [kJ per reaction]

Hydrogen releasing reactions

Ethanol and lactate


CH3COO + H+ + 2H2



CH3COO + HCO3  + H+ + 2H2


Fatty acids

 CH3COO + 4H2O

2HCO3  + H+ + 4H2



CH3COO + HCO3  + H+ + 3H2



2CH3COO + H+ + 2H2


 CH3(CH2)4COO + 3H2O

3CH3COO + 2H+ + 4H2



3CH3COO + 2H+ + H2


Glycolic acid

 HOCH2COO + H+ + H2O

2CO2 + 3H2


Aromatic compounds

 C6H5COO + 7H2O

3CH3COO + HCO3  + 3H+ + 3H2


 C6H5OH + 5H2O

3CH3COO + 3H+ + 2H2


 C6H4(OH)COO + 6H2O

3CH3COO + HCO3  + 3H+ + 2H2


Amino acids


CH3COO + HCO3  + NH4 + + H+ + 2H2

+ 7.5


CH3COO + 2HCO3  + NH4 + + H+ + 2H2



(CH3)2CHCH2COO + HCO3  + NH4 + + H+ + 2H2



CH3CH2COO + 2HCO3  + NH4 + + H+ + 2H2



CH3COO + 3HCO3  + 3H+ + 5H2


Hydrogen-consuming reactions

 4H2 + 2HCO3  + H+



 4H2 + HCO3  + H+

CH4 + 3H2O


 4H2 + 4So

4HS + 4H+


 4H2 + SO4 2− + H+

HS + 4H2O


 H2 + H2C(NH3 +)COO



  Open image in new window

  Open image in new window


 4H2 + NO3  + 2H+

NH4 + + 3H2O


aAll the calculations are based on published tables (Thauer et al. 1977; Kaiser and Hanselmann 1982; Dimroth 1983)

The final step involves two different groups of methanogens, the hydrogenotrophic methanogens which use the H2 and formate produced by other bacteria to reduce CO2 to CH4 and the acetotrophic methanogens which metabolize acetate to CO2 and CH4.

3 Microbiology biology of fatty acid degradation

Anaerobic degradation of LCFAs requires syntrophic communities of acetogenic bacteria, performing fatty acid β-oxidation and methanogenic archaea, which consume hydrogen and acetate to low concentrations (Schink 1997). In this syntrophic cooperation, interspecies hydrogen transfer plays a key role. Presently, about 14 species have been identified that are capable of degrading fatty acids in syntrophy with methanogens, all belonging to the families Syntrophomonadaceae within the group of low G + C-containing Gram-positive bacteria bacteria (McInerney 1992; Zhao et al. 1993; Wu et al. 2006; Sousa et al. 2007a) and Syntrophaceae in the subclass of the Deltaproteobacteria (Jackson et al. 1999). Of the 14 species identified so far, only seven are found capable of utilizing LCFAs with more than 12 carbon atoms, including Syntrophomonas sapovorans (Roy et al. 1986), Syntrophomonas saponavida (Lorowitz et al. 1989), Syntrophomonas curvata (Zhang et al. 2004), Syntrophomonas zehnderi (Sousa et al. 2007a), Syntrophomonas palmitatica (Hatamoto et al. 2007a), Thermosyntropha lipolytica (Svetlitshnyi et al. 1996) and Syntrophus aciditrophicus (Jackson et al. 1999). Among these microorganisms only four species have the capability of utilizing mono- and/or polyunsaturated LCFAs (with more than 12 carbon atoms): S. Sapovorans (Roy et al. 1986), S. curvata (Zhang et al. 2004), T. lipolytica (Svetlitshnyi et al. 1996) and the recently isolated S. zehnderi (Sousa et al. 2007a) (Table 6). The principle pathway of LCFAs degradation is via β-oxidation, but the initial steps involve in the conversion of unsaturated LCFAs are not yet fully understood. According to Weng and Jeris (1976) the degradation of LCFAs starts with the chain saturation, whereas other researchers provide evidence for direct β-oxidation of unsaturated LCFAs (Roy et al. 1986; Lalman and Bagley 2000, 2001). Generally, communities enriched on unsaturated LCFA also degrade saturated fatty acids, but the opposite does not always seem to be the case.
Table 6

Characteristics of some syntrophic LCFA-degrading bacteria (Source: Alves et al. 2009)

LCFA-degrading bacteria

Morphological characteristics

LCFA utilization range

Syntrophomonas sapovorans a

Short curved rods (0.5 × 2.5 μm)

Slightly motile


Two to four flagella

Non-spore forming

Degrades linear saturated fatty acids with 4–18 carbon atoms in co-culture with Methanospirillum hungatei. Mono- and di-unsaturated LCFA, such as oleate (C18:1) and linoleate (C18:2), are also oxidized by the co- culture

Syntrophomonas curvata b

Slightly curved rods (0.5–0.7 × 2.3–4.0 μm)



One or three flagella inserted in both poles

Non-spore forming

Degrades linear saturated fatty acids with 4–18 carbon atoms in co-culture with M. hungatei. Oleate (C18:1) is also oxidized by the co-culture

Syntrophus aciditrophicus c

Rod-shaped cells (0.5–0.7 × 1.0–1.6 μm)



Non-spore forming

Degrades linear saturated fatty acids with more than four carbon atoms (C4:0 to C8:0, C16:0, C18:0) in co-culture with H2-utilizingDesulfovibrio sp. or Methanospirillum hungatei

Syntrophomonas zehnderi d

Curved rods (approximately 0.4–0.7 ×

2.0–4.0 μm)

Variable response to Gram staining

Slight twitching


Spore formation during growth on oleate in

co-culture with a methanogen that utilizes hydrogen and formate

Degrades oleate, a mono-unsaturated fatty acid, and straight-chain fatty acids C4:0–C18:0 in syntrophic association with Methanobacterium formicicum

aRoy et al. (1986)

bZhang et al. (2004)

cJackson et al. (1999)

dSousa et al. (2007a)

In the last 20 years several biochemical and molecular methods have been employed to characterize the microbial communities involved in anaerobic degradation especially in anaerobic reactors such as CSTR, ABR, UASBR and EGSBR. The microbial diversity of anaerobic communities that are responsible for degrading LCFAs has not yet been studied extensively. But when the molecular diversity of granular and suspended sludge was assessed during a long term operation of two anaerobic up-flow bioreactors fed with the increasing concentrations of oleate, a physical segregation of the sludge in two fractions was observed. At the top there was a whitish floating fraction and a settled fraction at the bottom (Pereira et al. 2002). This segregation was due to the floatation caused by LCFAs accumulation. Differences between the structures of microbial communities of these two fractions were assessed by comparison of DGGE profiles. The values attained by the similarity indices between bottom and top sludge fractions for the granular and suspended sludge were as low as 56.7 and 29.4% respectively, while at the end of the operation the similarity indices between the original and suspended sludge, and the respective top sludge fractions were 17.3 and 15.2%.

Current developments in biological science using molecular approaches have widened the scope for studies on quantification and identification of the natural ecology of acetogenic bacteria (Hensen et al. 1999; Zhao et al. 1993). Hensen et al. (1999) used new oligonucleotide probes to characterize the phylogenic groups of mesophilic members of the family Syntrophomonadaceae as well as their syntrophic relationship with the hydrogenotrophic methanogens.

Shigematsu et al. (2006) used a 16S rRNA gene approach to study the microbial communities present in a chemostat fed with a mixture of oleic and palmitic acids. However, members belonging to Syntrophomonadaceae were detected in the chemostat, but the most predominant microorganisms identified, belonged to the Bacteroidetes and Spirochaetes phyla and it was suggested by authors that these members could also play a very important role in LCFAs degradation. Microbial diversity of anaerobic sludge after extended contact with long chain fatty acids (LCFAs) was studied, using molecular approaches. Sousa et al. (2007b, c) studied the diversity and dynamics of biomass in reactors treating saturated (palmitate, C16:0) and unsaturated (oleate, C18:1) LCFAs, using 16S ribosomal RNA genetargeted molecular techniques. During this study, it was found that the bacterial communities were dominated by the members of Clostridiaceae and Syntrophomonadaceae families (Sousa et al. 2007b). A new obligatory syntrophic bacterium, Syntrophomonas zehnderi, was isolated from an oleate-degrading culture and its presence in oleate-degrading sludges detected by 16S rRNA gene cloning and sequencing. Futhermore, presence of LCFA and short chain fatty acid degrading bacteria indicate that the formation of short chain fatty acids takes place by the degradation of LCFAs. However, a significant part of the retrieved bacterial 16S rRNA gene sequences (53%) were most similar to those of yet uncultured microorganisms, with the majority assigned to the phylum Firmicutes. Members of Proteobacteria and Bacteroidetes were also found in the culture.

Furthermore, it has been found that palmitate can be used by an oleate enrichment culture with no changes in the microbial community, while oleate cannot be used by the palmitate enrichment culture under the same condition; it suggests that palmitate is a key intermediate in oleate degradation. Communities enriched on unsaturated LCFAs also degrade saturated LCFAs, but the vice versa doesn’t occur in this case. The most likely biochemical degradation of unsaturated fatty acids starts with chain saturation (hydrogenation) followed by β-oxidation. Instead of Obligate hydrogen-producing acetogens (Schink 1997) that degrade unsaturated LCFAs, there is also the existence of microbial communities capable of hydrogenating unsaturated LCFAs to saturated LCFAs (Maia et al. 2007; Paillard et al. 2007). Stable isotope probing with 13C palmitate as a substrate was used by Hatamoto et al. (2007b, c) to find out the microorganisms directly involved in palmitate degradation. Members of the phyla Bacteroidetes and Spirochaetes, and the family Syntrophaceae within the Deltaproteobacteria, and members of the family Syntrophomonadaceae and genus Clostridium within the Firmicutes were found in clone libraries from heavy rRNA fractions. Thus, from these results it is evident that under methanogenic condition phylogenitically different groups of bacteria were active in situ in LCFA degradation. Anaerobic bacteria involved in the degradation of LCFAs in the presence of sulfate as electron acceptor, were studied by combined cultivation-dependent and molecular techniques (Sousa et al. 2009). Phylogenetic affiliation of rRNA gene sequences corresponding to predominant DGGE bands demonstrated that members of the Syntrophomonadaceae, together with sulfate reducers, mainly belonging to the Desulfovibrionales and Syntrophobacteraceae groups, were present in the sulfate-reducing enrichment cultures. The results of this study indicate that hydrogen consumption by methanogens is taken over by hydrogen-consuming sulfate reducers, which are known to have a higher affinity for hydrogen than methanogens.

4 Production of bioenergy and biochemicals from POME

Anthropogenic release of greenhouse gases, especially CO2 and CH4 has been recognized as one of the main causes of global warming. In Malaysia, the palm oil industry, particularly POME anaerobic treatment, has been identified as an important source of CH4. Due to increasing awareness of the risk of the environmental pollution and emission of green house gases (GHG), treatment of POME using biological processes in close digesters (UASBR) has gained popularity in the recent years, with over 500 installations worldwide (Latif et al. 2011a, b). POME is an ideal substrate for bioprocessing because it contains high level of degradable organic material which results in a net positive energy or economic balance. During the anaerobic treatment of POME, methane (Yacob et al. 2005, 2006a, b) and hydrogen (O-Thong et al. 2007) are generated, which can reduce the demand on energy resources and dependence on fossil fuels (O-Thong et al. 2008a).

5 Biological methane production

Methane has been categorized as one of the GHG with its global warming potential, 21 times more potent than CO2 (Ishigaki et al. 2005; Latif et al. 2011a, b). Methane fermentation offers an effective means of pollution reduction, superior to that achieved via conventional aerobic processes. Although practiced for decades, interest in anaerobic fermentation has only recently focused on its use in the economic recovery of fuel gas from industrial and agricultural surpluses. The anaerobic processes has considerable advantages over aerobic active sludge system such as (a) less energy demand (b) minimal sludge formation (c) minimization of unpleasant odour (d) efficient break down of organic substances by anaerobic bacteria to methane. In anaerobic digestion of POME, methanogenesis is the rate limiting step. Since anaerobic conventional digesters require large reactor and long retention time, therefore, high-rate anaerobic reactors have been proposed for POME treatment such as UASBR (Borja and Banks 1994a); upflow anaerobic filtration (Borja and Banks 1994b); fluidized bed reactor (Borja and Banks 1995a, b); and up-flow anaerobic sludge fixed-film reactor (Najafpour et al. 2006); anaerobic contact digester (Ibrahim et al. 1984) and continuous stirred tank reactor (Chin 1981), to reduce the reactor volume, shorten retention time along with the capture of methane gas. Table 7 lists the performance of various anaerobic treatment methods of POME.
Table 7

Performance of various anaerobic treatment methods on POME treatment (Source: Poh and Chong 2009)

Anaerobic treatment of palm oil mill effluent, practiced in Malaysia and Indonesia, results in domination of Methanosaeta concilii. It plays an important role in methane production from acetate and the optimum condition for its growth should be considered to harvest biogas as renewable fuel (Tabatabaei et al. 2009). Effective treatment of POME in UASBR is due to its ability to treat wastewater with high suspended solid contents (Fang and Chui 1994; Kalyunzai et al. 1998) that may clog reactors with packing material and also provide higher methane production (Kalyunzai et al. 1996; Stronach et al. 1978). However, this reactor might face long-startup periods if the seeded sludge is not granulated and this problem can be overcome by seeding the reactor with granulated sludge, resulting high performance along with shorter start-up period (Latif et al. 2011a, b).

Thus, anaerobic digestion process offers great potential for rapid disintegration of organic matter to produce methane that can be used to generate electricity (reaction 3) by burning it in a gas turbine or steam boiler (Linke 2006). Recovery of methane gas, from anaerobic treatment of POME as a renewable energy, represents a more acceptable alternative under the Kyoto Protocol with the objective of reducing the GHG emissions. Methane in the form of compressed natural gas is used as a vehicle fuel, and is claimed to be more environment friendly than other fossil fuels such as gasoline/petrol and diesel (reaction 2). In terms of energy it is comparable to commercially available gas fuels as shown in Table 8.
Table 8

Comparison between methane derived from anaerobic digestion of POME and other gas fuels

Chemical properties


Natural gas


Gas calorific value (K cal/kg)




Specific gravity




Ignition temperature (°C)




Inflammable limits (%)




Combustion air required (m3/m3)




Source: Ma (1999a, b)

Research is being conducted by NASA on methane’s potential as a rocket fuel (Lunar Engines, Aviation Week & Space Technology 2009). One advantage of methane is that it is abundant in many parts of the solar system and it could potentially be harvested in situ (i.e. on the surface of another solar-system body), providing fuel for a return journey (Barry 2007). The potential energy that could be generated from 1 m3 of biogas is 1.8 k Wh (Ma 1999a, b).

Recently methane emitted from coal mines has also been successfully converted to electricity and other useful products such as methanol for the use in the production of biodiesel e.g. by production of syngas (reaction 1 and 2) in downstream chemical processes (reaction 3). Syngas consists of H2 and carbon monoxide (CO) and very often some CO2, and has less than half the energy density of natural gas.
$$ {\text{CH}}_{ 4} + {\text{H}}_{ 2} {\text{O}} \rightleftarrows {\text{CO}} + 3 {\text{H}}_{ 2} $$
$$ {\text{CO}} + {\text{H}}_{ 2} {\text{O}} \rightleftarrows {\text{CO}}_{ 2} + {\text{H}}_{ 2} $$
$$ {\text{CH}}_{ 4} \left( {\text{g}} \right) + 2 {\text{O}}_{ 2} \left( {\text{g}} \right) \rightleftarrows {\text{CO}}_{ 2} \left( {\text{g}} \right) + 2 {\text{H}}_{ 2} {\text{O}}\left( {\text{l}} \right) + {\text{electricity}} $$

6 Biological hydrogen production

Biohydrogen is a promising energy carrier of the future: It is a promising clean fuel as it is ultimately derived from renewable energy sources, environment friendly (Ntaikou et al. 2010), since it burns to water (Guo et al. 2010; Mohammadi et al. 2011), gives high energy yield (142 MJ/kg), and can be produced by less energy-intensive processes (Nielsen et al. 2001; Das and Veziroglu 2001) and has a great potential to be an alternative fuel. Cellulose and starch containing biomass can be used as a reliable and renewable raw material for hydrogen gas production. Due to the nature of POME, with high cellulose and lignocellulosic material, it takes a long time to degrade the organic substances and has a potential as a substrate for generation of hydrogen (Fakhru’l-Razi et al. 2005; Vijayaraghavan and Ahmad 2006). Biohydrogen fermentation can be realized by three main categories: (a) Dark fermentation (b) Photo fermentation (c) dark-photo fermentation (Manish and Banerjee 2008; Tao et al. 2007).

Research in dark fermentation, for hydrogen production, is on the increase in recent years due to its potential importance in our economy (Logan 2002; Yokoi 2002; Yu 2002; Zhang 2003) and has presented a promising route of biohydrogen production compared to photosynthetic routes (Levin et al. 2004). The major advantages of dark fermentation are high rate of cell growth, no light energy requirement, no oxygen limitation problems and ability to run on low capital cost (Levin et al. 2004; Nath and Das 2004; Hallenbeck and Benemann 2002). A variety of bacteria such as E. coli, Enterobacter aerogenes, and Clostridium butyricum are known to ferment sugars and produce hydrogen, using multienzyme systems. Since these “dark fermentation” reactions do not require light energy, so they are capable of constantly producing hydrogen from organic compounds throughout the day and night. Compared with other biological hydrogen-production processes, fermentative bacteria have high evolution rates of hydrogen. However, sugars are relatively expensive substrates that are not available in sufficient quantities to support hydrogen production at a scale required to meet energy demand. Photosynthetic hydrogen production basically involves two pathways. (1) Algae (Cyanobacteria) break down water to H2 and O2 in the presence of light energy. (2) Photo-heterotrophic bacteria ((Rhodobacter sp.) utilize organic acids (98–99%) such as acetic, lactic and butyric acids to produce H2 (Tao et al. 2007). Photo-fermentation requires light in the ultra-violet range up to 400 nm (Kahn and Durako 2009) and the UV light has the potential to eliminate foreign micro-organisms and to prevent contamination but the production rates with photo-fermentation is not as high as with dark-fermentation. However, various researchers reported that anaerobic microflora found in POME was able to produce hydrogen (Chong et al. 2009a, b, c) and methane gas was not observed in the evolved gas (Morimoto et al. 2004; Atif et al. 2005). Vijayaraghavan and Ahmad (2006) isolated hydrogen generating microflora from the cow dung in POME, based on pH adjustment (pH 5) coupled with heat treatment proved to be promising candidate towards hydrogen generation. Hydrogen production and growth of hydrogen producing bacteria were increased by nutrient (N, P and Fe) supplementation (O-Thong et al. 2007, 2008a, b). Furthermore, from the Table 9, it is evident that dark- fermentation can produce hydrogen more efficiently than photo-fermentation. The major advantages of the 2nd pathway of photo-fermentation is more favorable process economy, reduced operating problems and higher rate of H2 gas production. Organic acids produced in the acidogenic phase of anaerobic digestion of organic materials may be used as the substrate for production of hydrogen by the photo-heterotrophic organisms. Therefore, the hydrogen yield may be improved by using a two-stage process such as dark and photo-fermentations or by their combinations. Hydrolysis (acid or enzymatic) of starch/cellulose to highly concentrated sugar solution is the first step in fermentative hydrogen production from waste biomass followed by dark fermentation of resulting carbohydrates to volatile fatty acids (VFA), H2 and CO2 by acetogenic-anaerobic bacteria. Usually photo-heterotrophic bacteria (Rhodobacter sp.) are used for photo-fermentation of VFAs to CO2 and H2 (Kapdan and Kargi 2006; Manish and Banerjee 2008). Fang et al. (2005) reported that a maximum hydrogen yield, 2.5 mol-H2/mol-acetate and 3.7 mol-H2/mol-butyrate could be achieved by Rhodobacter capsulatus. Thus, the effluents of dark-fermentation, which are mainly composed of organic acids, could be used as the substrate of photosynthetic non sulphur bacteria to produce hydrogen. Redwood and Macaskie (2006), used glucose and Tao et al. (2007) used sucrose as the substrate in the two-step fermentation process, and found remarkably improved compared with that of dark-fermentation. A combined biohydrogen and biomethane generation process has been proposed for organic solid wastes (Ueno et al. 2007), food wastes (Han and Shin 2004), cheese whey (Antonopoulou et al. 2008), olive mill wastewater (Koutrouli et al. 2009) and wastewater sludge (Ting and Lee 2007). As for as POME is concerned, neither dark-photofermentation nor dark-methanogenic process has yet been investigated. Therefore, biological and engineering studies need to concentrate on these issues for commercial viability and sustainable energy production from POME.
Table 9

Yield of biohydrogen production from POME using dark and photo-fermentation technique




Temperature (°C)

Biohydrogen yield (L H2/L POME)


Dark fermentation


Mixed culture

uncont rolled


23.82 mmol H2/(l-medium)

Morimoto et al. (2004)


Mixed culture



4.708 ml H2/(l-medium)

Atif et al. (2005)


Mixed culture


0.42 l biogas/g COD destroyed with 57% hydrogen content

Vijayaraghavan and Ahmad (2006)





4.4 l H2/(l-medium) per day

O-Thong et al. (2007)





6.1 l H2/(l-medium per day)

O-Thong et al. (2007)






O-Thong et al. (2008a, b)


Clostridium butyricum




Chong et al. (2009b)


Clostridium butyricum




Chong et al. (2009c)


Mixed culture




Yusoff et al. (2009)



Rhodopseudomonas palustris



Jamil et al. (2009)

7 Production of bioplastics (PHA)

PHA (Fig. 2) is a group of biodegradable polymers, with properties similar to polypropylene and polyethylene (Salehizadeh and Van Loosdrecht 2004). Unlike normal plastics, which are non-biodegradable, these bioplastics biodegrade naturally to carbon dioxide and water within a few weeks upon disposal in the soil or the environment. PHA is an energy and carbon storage molecule produced in bacteria during growth limiting conditions (Lee 1996; Salehizadeh and Van Loosdrecht 2004). Due to the problems and harmful effects of conventional plastics on the environment, there has been a considerable interest in the development of biodegradable plastics for the last few decades (Lee and Yu 1997; Chua et al. 2003; Kumar et al. 2004). PHB and its copolymer, poly (3-hydroxybutyrate-co-3-hydroxyvalerate) [P(3HB-co-3-HV)] are the most widespread PHAs, although other forms are also possible. However, the current pure culture production is expensive. The high cost of PHA has so far caused it to play a marginal role in the world’s plastic market [120]. To decrease the production cost of these biopolymers, open mixed cultures can be used for PHA production (Albuquerque et al. 2007). Several workers have studied PHA production by mixed cultures using synthetic organic acids (Dionisi et al. 2005a, b; Lemos et al. 2006; Bengtsson et al. 2008). In recent years, to reduce the cost price considerably, POME has gained great attention by the research institutions and industrial sectors due to its potential as sources of carbon and nitrogen for microbial growth (Hassan et al. 1997a, b, c, 2002; Zakaria et al. 2008). A suggested three step process for PHA production in open mixed cultures, consists of: (1) acidogenic fermentation of the waste stream to produce VFAs, (2) selection of PHA storing organisms, (3) accumulation of PHA in the selected biomass (Dionisi et al. 2005b).
Fig. 2

(i) Structure of PHA. (ii) Metabolic pathway for Medium Chain Length (MCL)-PHA synthesis from fatty acid with confirmed or postulated enzymes (Source: Van der Leij and Witholt 1995)

Since, most prokaryotes are capable of PHA production (Chua et al. 2003), therefore, it was possible to use the mixed cultures to produce PHA in POME (Din et al. 2006a, b). The initial mixed culture was developed by using 10% activated sludge from the sewage treatment plant and 90% from palm oil mill effluent (POME). It was observed that by using mixed cultures and POME, different kinds of PHA-constituent could be obtained and its harvesting was more reliable for the use of biodegradable plastics than a single PHA-constituent. The cultivation was maintained in a single batch-fed reactor and operated in two steps: First the system was allowed for an extensive growth (using a nutrient medium), and then a feed with limited nutrients (no nutrient medium adaptation) will be introduced in the next step. The average production of PHA, by using POME, could only reach up to 44% of the cell by dry weight. However, the favourable factors (e.g. temperature and harvesting time) have been made in the next stage to induce the PHA production (Din et al. 2006a, b). Table 10 shows the production of PHA from POME.
Table 10

PHA production from POME



Temp (°C)



Maximum production g/l



Rhodobactor spheroids

IFO 12203





Hassan et al. (1996)

Rhodobactor spheroids

IFO 12203





Hassan et al. (1997b)

Alcaligenes eutrophus H 16 (ATCC 17699)



Stirred tank


Hassan et al. (1997c)

Ralstonia eutropha

ATCC 17699



Bioreactor fermentati-on


Hassan et al. (2002)

Mixed cultures



Sequencin-g batch reactor


Md Din et al. (2006b)

A new isolate, designated as strain EB172, was isolated from a digester tank, treating palm oil mill effluent and was investigated by polyphasic taxonomic approaches (Zakaria et al. 2010). The cells were rod-shaped, Gram-negative, non-pigmented, non-spore-forming and non-fermentative. Biochemical test, microscopic observation, cellular fatty acids analysis and 16S rRNA gene sequence analyses revealed that this strain was placed in the cluster of genus Comamonas and differed from the existing Comamonas species. Furthermore, the strain was capable of accumulating PHA up to 59% of CDW in fed-batch cultivation process with pH–stat continuous feeding of mixed organic acids derived from anaerobically treated POME (Zakaria et al. 2010).

Since POME contains high organic content, almost being non-toxic, can be suitable for PHA production. VFAs can be stored as PHA by bacteria in the opened mixed cultures. For the selection of PHA storing organisms, the feast-famine approach is most commonly used (Dionisi et al. 2005a; Albuquerque et al. 2007; Bengtsson et al. 2008). In the feast-famine selection, the substrate is fed in pulses, whose concentration alternates from high (feast) to low (famine) (Dionisi et al. 2004). A final accumulation step is used to saturate the biomass with PHA (Dionisi et al. 2004).

8 Production of organic acids

Besides, biogas/methane generation from anaerobic treatment of POME [64], that can also be used to generate organic acids, which are effectively used in a variety of industrial processes (Hassan et al. 2002; Mumtaz et al. 2008). During anaerobic treatment of POME, the organic acids are produced as an intermediate which can be used as raw material for PHA production (Hassan et al. 1996, 1997b, c, 2002). Biotransformation of hexoses to gluconic, itaconic, citric and lactic acids is being carried out on a large scale to produce basic ingredients for laundry detergents, glues, preservatives and polylactides. Plastic materials, made from polylactides, are used in medicines related to transplantations, due to its excellent compatibility with tissues and skin. The production of mixed organic acids, from anaerobicallay treated palm oil mill effluent, has been shown to be a renewable and cheaper carbon sources of PHAs production (Hassan et al. 1997a, b, c, 2002; Zakaria et al. 2008). Hassan et al. (1996) reported that more than 70% of the BOD sources in POME, were converted to organic acids by R. spheroids IFO 12203 and a 50% PHAs yield from organic acids achieved. The predominant organic acids, at a higher and lower pH, were acetic acid and formic acid respectively. Freezing and thawing of treated POME was successfully applied to separate sludge solids in conjunction with sludge recycle system, so that continuous anaerobic treatment retention time can be shortened to 3.5 days (Phang et al. 2002, 2003). The performance of the anaerobic treatment of palm oil mill effluent for the production of organic acids at a short retention time of less than 5 days, was carried out by Lee et al. (2003) by incorporating a sludge recycle system without pH control and calcium carbonate. They showed that by incorporating a sludge recycle system with the freezing-thawing method, the retention time for the treatment could be reduced to 3.5 days without affecting the generation of organic acids.

The production of itaconic acid by Aspergillus terreus IMI 282743 from filtered Palm Oil Mill Effluent (POME), as an added supplement, was investigated by Jamaliah et al. Jamaliah et al. (2006). Itaconic acid (methylene butanedioic acid; common synonyms: methylene succinic acid, 3-carboxy-3-butanoic acid, propylenedicarboxylic acid) is one of the promising substances within the group of organic acids. Jamal et al. (2005) used POME and wheat flour as a medium and screened potential microorganisms for citric acid production and showed that Aspergillus (A 103) produced the highest concentration of citric acid after 2 days of fermentation.

Alam et al. (2008) investigated the bioconversion of POME, by adding co-substrate (glucose and wheat flour and nitrogen source–ammonium nitrate), for citric acid production under optimal conditions and observed higher removal of chemical oxygen demand (82%) with the production of citric acid (5.2 g/l) on the final day of fermentation process (7 days). Mumtaz et al. (2008) generated organic acids with low molecular weight such as acetic acid, propionic acid and butyric acids from partial anaerobic treatment of palm oil mill effluent (POME), using pilot scale filtration and evaporation system. The recovery of organic acids has a significant and economical impact, since around 50% cost of PHA production is believed to be associated with the substrate itself. Thus, the organic acid generated during acid-phase anaerobic digestion of POME can be converted into value-added products and fine chemicals and may serve as a renewable feed stock for biosynthesis of PHA. Table 11 summarizes production of organic acid from POME.
Table 11

Organic acid produced from POME




Temp (°C)


Fermentation time

Maximum production g/l


Organic acids

Mixed cultures



Bioreactor fermentation



Hassan et al. (1996)

Organic acids

Mixed cultures



Stirred tank



Yee et al. (2003)

Citric acid

Aspergillus (A103)



Flask fermentation



Jamal et al. (2005)

Citric acid

Aspergillus niger (A103)



Flask fermentation



Alam et al. (2008)

Itaconic acid

Aspergillus terreus IMI 282743


Flask fermentation



Wu et al. (2005)

9 Conclusion

Renewable energy has been identified globally as a key driver to achieve economic growth while ensuring minimal environmental harm. Simultaneously, the current development of green technology and its related policies have enhanced the growth of renewable energy in the country.

The use of POME as a renewable energy resource can improve energy security while reducing the environmental burdens of waste disposal. The Malaysian palm oil industry, with 4.69 million hectares of planted land has a tremendous opportunity in supplying renewable energy. Energy from wastewater therefore facilitates the integration of water, waste and energy management within a model of sustainable development. Presently, renewable energy represents 5% of all prime energy use, but by the year 2060, it is strongly predicted that it will reach 70%. It is estimated that these palm based materials could generate up to 1,260 MW of energy. This amounts to nearly 10% of the maximum energy demand of electricity in Malaysia. Therefore, oil palm-based biomass can be expected to play a prominent role in the future when the demand for renewable energy is expected to increase rapidly.

In Malaysia, 5% of the electricity generated by renewable energy can easily be met by renewable sources. POME can be treated anaerobically to breakdown organic matters while releasing biomethane and sometimes, biohydrogen. Lipids are suitable substrates for high-rate anaerobic wastewater treatment and are also ideal co-substrates for AD plants. By utilizing appropriate technology and following the right feeding strategy, lipids can be effectively converted to methane by syntrophic consortia of acetogenic bacteria and methanogenic archaea. Application of cultivation and molecular techniques to the study of microbial composition of LCFA-degrading sludges provided important insight into the communities involved in the degradation of these compounds.



The author would like to thank Faculty of Civil Engineering and Earth Resources and R & D Cluster unit for technical assistance during the course of this research. Moreover, the financial support from the Fundamental Research Grant Scheme (RDU-0903113, University Malaysia Pahang) is highly acknowledged and appreciated.

Open Access

This article is distributed under the terms of the Creative Commons Attribution Noncommercial License which permits any noncommercial use, distribution, and reproduction in any medium, provided the original author(s) and source are credited.


  1. Agamuthu P, Tan EL (1985) Digestion of dried palm oil mill effluent by Cellulomonas sp. Microbiol Lett 30:109–113Google Scholar
  2. Ahmad AD, Ismail S, Bhatia S (2003) Water recycling from palm oil mill effluent (POME) using membrane technology. Desalination 157:87–95CrossRefGoogle Scholar
  3. Alam MZ, Jamal P, Nadzir MM (2008) Bioconversion of palm oil mill effluent for citric acid production: statistical optimization of fermentation media and time by central composite design. World J Microbiol Biotechnol 24:1177–1185CrossRefGoogle Scholar
  4. Albuquerque MGE, Eiroa M, Torres C, Nunes BR, Reis MAM (2007) Strategies for the development of a side stream process for polyhydroxyalkanoate (PHA) production from sugar cane molasses. J Biotechnol 130:411–421CrossRefGoogle Scholar
  5. Alves MM, Pereira MA, Sousa DZ, Cavaleiro AJ, Picavet M, Smidt H, Stams AJM (2009) Waste lipids to energy: how to optimize methane production from long-chain fatty acids (LCFA). Microbial Biotechnol 2(5):538–550CrossRefGoogle Scholar
  6. Andreasen T (1982) The AMINODAN system for treatment of palm oil mill effluent. In: Proceeding of regional workshop on palm oil mill technology and effluent treatment. PORIM, Malaysia, pp 213–215Google Scholar
  7. Angelidaki I, Ahring BK (1995) Establishment and characterization of anaerobic thermophilic (55°C) enrichment culture degrading long chain fatty acids. App Environ Microbiol 61(6):2442–2445Google Scholar
  8. Antonopoulou G, Stamatelatou K, Venetsaneas N, Kornaros M, Lyberatos G (2008) Biohydrogen and methane production from cheese whey in a two-stage anaerobic process. Ind Eng Chem Res 47:5227–5233CrossRefGoogle Scholar
  9. Atif AAY, Fakhru’a-Razi A, Ngan MA, Morimoto M, Iyuke SE, Vesiroglu NT (2005) Fed batch production of hydrogen from palm oil mill effluent using anaerobic microflora. Int J Hydrogen Energy 30:1393–1397CrossRefGoogle Scholar
  10. Barry P (2007) Methane Blast. NASA. May 4, 2007Google Scholar
  11. Basri MF, Yacob S, Hassan MA, Shirai Y, Wakisaka M, Zakaria MR, Phang LY (2010) Improved biogas production from palm oil mill effluent by a scaled-down anaerobic treatment process. World J Microbiol Biotechnol 26:505–514CrossRefGoogle Scholar
  12. Batstone DJ (2000) High rate anaerobic treatment of complex waste water. PhD Thesis, Queensland University, AustraliaGoogle Scholar
  13. Beccari M, Bonemazzi F, Majone M, Reccardi C (1996) Interaction between acidogenesis and methanogenesis in the anaerobic treatment of olive oil mill effluents. Water Res 30(1):183–189CrossRefGoogle Scholar
  14. Bengtsson S, Werker A, Christensson M, Welander T (2008) Production of polyhydroxyalkanoates by activated sludge treating a paper mill wastewater. Bioresour Technol 99:509–516CrossRefGoogle Scholar
  15. Biebl H, Pfening N (1978) Growth yields of green sulfur bacteria in mixed cultures with sulfur and sulfate reducing bacteria. Arch Microbiol 117:9–16CrossRefGoogle Scholar
  16. Bitton B (2005) Wastewater microbiology, 3rd Ed. Wiley, Hoboken, New Jersey, Chap 13, 2005 pp 345–369Google Scholar
  17. Borja R, Banks CJ (1994a) Anaerobic digestion of palm oil mill effluent using an up-flow anaerobic sludge blanket (UASB) reactor. Biomass Bio 6:381–389CrossRefGoogle Scholar
  18. Borja R, Banks CJ (1994b) Kinetic of methane production from palm oil mill effluent in an immobilized cell bioreactor using saponite as support medium. Biores Technol 48:209–214CrossRefGoogle Scholar
  19. Borja R, Banks CJ (1994c) Treatment of palm oil mill effluent by upflow anaerobic filtration. J Chem Technol Biotechnol 61:103–109CrossRefGoogle Scholar
  20. Borja R, Banks CJ (1995a) Response of an anaerobic fluidized bed reactor treating ice-cream wastewater to organic, hydraulic, temperature and pH shocks. J Biotechnol 39:251–259CrossRefGoogle Scholar
  21. Borja R, Banks CJ (1995b) Comparison of an anaerobic filter and an anaerobic fluidized bed reactor treating palm oil mill effluent. Process Biochem 30:511–521Google Scholar
  22. Chan KS, Chooi CF (1982) Ponding system for palm oil mill effluent. In: PORIM proceedings of regional on palm oil mill technology and effluent treatment, pp 185–192Google Scholar
  23. Cheng J, Zhu X, Borthwick A (2010) Palm oil mill effluent treatment using a two-stage microbial fuel cells system integrated with immobilized biological aerated filters. Bioresour Technol 101:2729–2734CrossRefGoogle Scholar
  24. Chin KK (1981) Anaerobic treatment kinetics of palm oil sludge. Water Res 15:199–202CrossRefGoogle Scholar
  25. Chong ML, Sabaratnam V, Shirai Y, Hassan MA (2009a) Biohydrogen production from biomass and industrial wastes by dark fermentation. Int J Hydrogen Energy 34:3277–3287CrossRefGoogle Scholar
  26. Chong ML, Abdul Rahman NA, Rahim RA, Aziz SA, Shirai Y, Hassan MA (2009b) Optimization of biohydrogen production by Clostridium butyricum EB6 from palm oil mill effluent using response surface methodology. Int J Hydrogen Energy 34:7475–7482CrossRefGoogle Scholar
  27. Chong ML, Rahim RA, Shirai Y, Hassan MA (2009c) Biohydrogen production by Clostridium butyricum EB6 from palm oil mill effluent. Int J Hydrogen Energy 34:764–771CrossRefGoogle Scholar
  28. Chua ASM, Takabatake H, Satoh H, Mino T (2003) Production of polyhydroxyalkanoates (PHA) by activated sludge treating municipal wastewater: effect of pH, sludge retention time (SRT) and acetate concentration in influent. Water Res 37:3602–3611CrossRefGoogle Scholar
  29. Das D, Veziroglu TN (2001) Hydrogen production by biological processes: a survey of literature. Int J Hydrogen Energy 26:13–28CrossRefGoogle Scholar
  30. Demirel B, Scherer P (2008) The roles of acetotrophic and hydrogenotrophic methanogens during anaerobic conversion of biomass to methane a review. Rev Environ Sci Biotechnol 7:173–190CrossRefGoogle Scholar
  31. Dimroth K (1983) Thermochemische daten organischer verbindungen. In D’ans-Lax Taschenbuch für Chemiker und Physiker, Bd. 2. Springer, Berlin, pp 997–1083Google Scholar
  32. Din MF, Ujang Z, Muhd Yunus S, Van Loosdrecht MCM (2006) Storage of polyhydroxyalkanoates (PHA) in fed-batch mixed cultures. Fourth Seminar on Water Management (JSPS-VCC). Johor, Malaysia July 11–13Google Scholar
  33. Din MF, Ujang Z, van Loosdrecht MCM, Ahmad A, Sairan MF (2006b) Optimization of nitrogen and phosphorus limitation for better biodegradable plastic production and organic removal using single fed-batch mixed cultures and renewable resources. Water Sci Technol 53:15–20CrossRefGoogle Scholar
  34. Dionisi D, Majone M, Papa V, Beccari M (2004) Biodegradable polymers from organic acids by using activated sludge enriched by aerobic periodic feeding. Biotech Bioeng 85(6):569–579CrossRefGoogle Scholar
  35. Dionisi D, Beccari M, Di Gregorio S, Majone M, Petrangeli Papini M, Vallini G (2005a) Storage of biodegradable polymers by an enriched microbial community in a sequencing batch reactor operated at high organic load rate. J Chem Technol Biotechnol 80:1306–1318CrossRefGoogle Scholar
  36. Dionisi D, Carucci G, Papini MP, Riccardi C, Majone M, Carrasco F (2005b) Olive oil mill effluents as a feedstock for production of biodegradable polymers. Water Res 39:2076–2084CrossRefGoogle Scholar
  37. Faisel M, Unno H (2001) Kinetic analysis of palm oil mill wastewater treatment by a modified anaerobic baffled reactor. Biochem Eng J 9:25–31CrossRefGoogle Scholar
  38. Fakhru’l-Razi A, Yassin AAA, Lyuke SE, Ngan MA, Morimoto M (2005) Biohydrogen synthesis from wastewater by anaerobic fermentation using microflora. Int J Green Energy 2:387–396CrossRefGoogle Scholar
  39. Fang HHP, Chui HK (1994) Comparison of startup performance of four anaerobic reactors for the treatment of high-strength wastewater. Resour Conserv Recycling 11:123–138CrossRefGoogle Scholar
  40. Fang HHP, Liu H, Zhang T (2005) Phototrophic hydrogen production from acetate and butyrate in wastewater. Int J Hydrogen Energy 30(7):785–793CrossRefGoogle Scholar
  41. Federal Subsidiary Legislation (1974) Federal subsidiary legislation: environmental quality act 1974 [ACT 127], environmental quality (Sewage and Industrial Effluents) regulation 1979. Available: (29/2/2008)
  42. Foo KY, Hameed BH (2010) Insight into the applications of palm oil mill effluent: a renewable utilization of the industrial agricultural waste. Renew Sustain Energy Rev 14:1445–1452CrossRefGoogle Scholar
  43. Guo XM, Trably E, Latrille E, Carrère H, Steyer JP (2010) Hydrogen production from agricultural waste by dark fermentation: a review. Int J Hydrogen Energy doi:  10.1016/j.ijhydene.2010.03.008
  44. Habib MAB, Yusoff FM, Phang SM, Ang KJ, Mohamed S (1997) Nutritional values of chironomid larvae grown in palm oil mill effluent and algal culture. Aquaculture 158:95–105CrossRefGoogle Scholar
  45. Hallenbeck PC, Benemann JR (2002) Biological hydrogen production: fundamentals and limiting processes. Int J Hydrogen Energy 27:1185–1193CrossRefGoogle Scholar
  46. Han SK, Shin HS (2004) Performance of an innovative two-stage process converting food waste to hydrogen and methane. J Air Waste Manage Assoc 54:242–249Google Scholar
  47. Hanaki K, Nagase M, Matsuo T (1981) Mechanism of inhibition caused by long-chain fatty acids in anaerobic digestion process. Biotechnol Bioeng 23:1591–1610CrossRefGoogle Scholar
  48. Hanaki K, Matsuo T, Nagase M, Tabata Y (1987) Evaluation of effectiveness of two-phase anaerobic digestion process degrading complex substrate. Water Sci Technol 19:311–322Google Scholar
  49. Hassan MA, Shirai Y, Kusubayashi N, Karim MIA, Nakanishi K, Hashimoto K (1996) Effect of organic acid profiles during anaerobic treatment of palm oil mill effluent on the production of polyhydroxyalkanoates by Rhodobacter sphaeroides. J Ferment Bioeng 82:151–156CrossRefGoogle Scholar
  50. Hassan MA, Shirai Y, Umeki H, Yamazumi H, Jin S, Yamamoto S, Abdul Karim MI, Nakanishi I, Hashimoto K (1997a) Acetic acid separation from anaerobically treated pal oil mill effluent by ion exchange resins for the production of polyhydroxyalkanoate by Alcaligens eutrophus. Biosci Biotechnol Biochem 61(9):1465–1468CrossRefGoogle Scholar
  51. Hassan MA, Shirai Y, Kusubayashi N, Karim MIA, Nakanishi K, Hashimoto K (1997b) The production of polyhydroxyalkanoate from anaerobically treated palm oil mill effluent by Rhodobacter sphaeroides. J Ferment Bioeng 83:485–488CrossRefGoogle Scholar
  52. Hassan MA, Shirai Y, Umeki H, Yamazumi H, Jin S, Yamamoto S (1997c) Acetic acid separation from anaerobically treated palm oil mill effluent by ion exchange resins for the production of polyhydroxyalkanoate by Alcaligenes eutrophus. Biosci Biotechnol Biochem 61:1465–1468CrossRefGoogle Scholar
  53. Hassan MA, Nawata O, Shirai Y, Rahman NAA, Yee PL, Ariff A (2002) A proposal for zero emission from palm oil industry incorporating the production of polyhydroxyalkanoates from palm oil mill effluent. J Chem Eng Jpn 35:9–14CrossRefGoogle Scholar
  54. Hatamoto M, Imachi H, Yashiro Y, Fukayo S, Ohashi A, Harada H (2007a) Syntrophomonas palmitatica sp nov., an anaerobic, syntrophic, long-chain fatty-acidoxidizing bacterium isolated from methanogenic sludge. Int J Syst Evol Microbiol 57:2137–2142CrossRefGoogle Scholar
  55. Hatamoto M, Imachi H, Yashiro Y, Ohashi A, Harada H (2007b) Diversity of anaerobic microorganisms involved in long-chain fatty acid degradation in methanogenic sludges as revealed by RNA-based stable isotope probing. Appl Environ Microbiol 73:4119–4127CrossRefGoogle Scholar
  56. Hatamoto M, Imachi H, Ohashi A, Harada H (2007c) Identification and cultivation of anaerobic, syntrophic longchain fatty acid-degrading microbes from mesophilic and thermophilic methanogenic sludges. Appl Environ Microbiol 73:1332–1340CrossRefGoogle Scholar
  57. Hensen KH, Ahring BK, Raskin L (1999) Quantification of syntrophic fatty acid-beta-oxidizing bacteria in a mesophilic biogas reactor by oligonucleotide probe hybridization. Appl Environ Microbiol 65:4767–4774Google Scholar
  58. Ibrahim A, Yeoh BG, Cheah SC, Ma AN, Ahmad S, Chew TY, Raj R, Wahid MJA (1984) Thermophilic anaerobic contact digestion of palm oil mill effluent. Water Sci Technol 17:155–166Google Scholar
  59. Ishigaki T, Yamada M, Nagamori M, Ono Y, Inoue Y (2005) Estimation of methane emission from whole waste landfill site using correlation between flux and ground temperature. Environ Geol 48:845–853CrossRefGoogle Scholar
  60. Jackson BE, Bhupathiraju VK, Tanner RS, Woese CR, McInerney MJ (1999) Syntrophus aciditrophicus sp. nov., a new anaerobic bacterium that degrades fatty acids and benzoate in syntrophic association with hydrogen-using microorganisms. Arch Microbiol 171:107–114CrossRefGoogle Scholar
  61. Jamal P, Alam MZ, Salleh MRM, Nadzir MM (2005) Screening of microorganisms for citric acid production from palm oil mill effluent. Biotechnology 4:275–278CrossRefGoogle Scholar
  62. Jamaliah MJ, Muhammad NIS, Yeong WT (2006) Factor analysis in itaconic acid fermentation using filtered POME by Aspergillus terreus IMI 282743. J Kejurutraan 18:39–48Google Scholar
  63. Jamil Z, Mohamad Annuar MS, Ibrahim S, Vikineswary S (2009) Optimization of phototrophic hydrogen production by Rhodopseudomonas palustris PBUM001 via statistical experimental design. Int J Hydrogen Energy 34:7502–7512CrossRefGoogle Scholar
  64. Jeris JS, McCarty M (1965) The biochemistry of methane fermentation using 14C tracers. J Water Pollut Control Fed 37:178–192Google Scholar
  65. Kahn AE, Durako MJ (2009) Wavelength-specific photo-synthetic responses of Halophila johnsonii from marine-influenced versus river-influenced habitats. Aquat Bot 91(3):245–249CrossRefGoogle Scholar
  66. Kaiser JP, Hanselmann KW (1982) Fermentative metabolism of substituted monoaromatic compounds by a bacterial community from anaerobic sediments. Arch Microbiol 133:185–194CrossRefGoogle Scholar
  67. Kalyunzai SV, Skylar VI, Davlyatshina MA, Parshina SN, Simankova MV, Kostrikina NA, Nozhevnikova AN (1996) Organic removal and microbiological features of UASB-reactor under various organic loading rates. Biores Technol 55:47–54CrossRefGoogle Scholar
  68. Kalyunzai S, de los Santos LE, Martinez JR (1998) Anaerobic treatment of raw and preclarified potato-maize wastewater in a UASB reactor. Biores Technol 66:198–199Google Scholar
  69. Kapdan IK, Kargi F (2006) Biohydrogen production from waste materials. Enzym Microb Technol 38:569–582CrossRefGoogle Scholar
  70. Kim SH, Han SK, Shin HS (2004) Two-phase anaerobic treatment system for fat-containing wastewater. J Chem Technol Biotechnol 79:63–71CrossRefGoogle Scholar
  71. Komatsu T, Hanaki K, Matsuo T (1991) Prevention of lipid inhibition in anaerobic processes by introducing a two-phase system. Water Sci Technol 23:1189–1200Google Scholar
  72. Koutrouli EC, Kalfas H, Gavala HN, Skiadas IV, Stamatelatou K, Lyberatos G (2009) Hydrogen and methane production through two-stage mesophilic anaerobic digestion of olive pulp. Bioresour Technol 100:3718–3723CrossRefGoogle Scholar
  73. Kumar S, Tamura K, Nei M (2004) MEGA 30: Integrated software of molecular evolutionary genetics analysis and sequence alignment. Briefings in Bioinformatics 5(2):150–163CrossRefGoogle Scholar
  74. Lalman JA (2000) Anaerobic degradation of linoleic (C18:2), oleic (C18:1) and stearic (C18:0) acids and their inhibitory effects on acidogens, acetogens and methanogens. Ph.D. thesis, University of Toronto: Toronto, CanadaGoogle Scholar
  75. Lalman JA, Bagley DM (2000) Anaerobic degradation and inhibitory effects of linoleic acid. Water Res 34:4220–4228CrossRefGoogle Scholar
  76. Lalman JA, Bagley DM (2001) Anaerobic degradation and methanogenic inhibitory effects of oleic and stearic acids. Water Res 35:2975–2983CrossRefGoogle Scholar
  77. Latif MA, Ahmad A, Ghufran R, Wahid ZA, (2011) Effect of temperature and organic loading rate on upflow anaerobic sludge blanket reactor and CH4 production by treating liquidized food waste. Environmental Prog & Sustain Ener doi: 10.1002/ep.10540
  78. Latif MA, Ghufran R, Wahid ZA, Ahmad A (2011) Integrated application of upflow anaerobic sludge blanket reactor for the treatment of wastewaters. Water Res 05–049Google Scholar
  79. Lee SY (1996) Bacterial polyhydroxyalkanoates. Biotechnol Bioeng 49:1–14CrossRefGoogle Scholar
  80. Lee S, Yu J (1997) Production of biodegradable thermoplastics from municipal sludge by two-stage bioprocess. Resour Conserv Recycling 19:151–164CrossRefGoogle Scholar
  81. Lee HJ, Lee SC, Kim JD, Oh YG, Kim BK, Kim CW, Kim KJ (2003) Methane production potential of feed ingredients as measured by in vitro gas test. Asian-Aust J Anim Sci 16(8):1143–1150Google Scholar
  82. Lemos PC, Serafim LS, Reis MAM (2006) Synthesis of polyhydroxyalkanoates from different short-chain fatty acids by mixed cultures submitted to aerobic dynamic feeding. J Biotechnol 122:226–238CrossRefGoogle Scholar
  83. Levin DB, Pitt L, Love M (2004) Biohydrogen production: prospects and limitations to practical application. Int J Hydrogen Energy 29:173–185CrossRefGoogle Scholar
  84. Linke B (2006) Kinetic study of thermophilic anaerobic digestion of solid wastes from potato processing. Biomass Bioenergy 30:892–896CrossRefGoogle Scholar
  85. Logan BE (2002) Biological hydrogen production measured in batch anaerobic respirometers. Environ Sci Technol 36:2530–2535CrossRefGoogle Scholar
  86. Lorowitz WH, Zhao HX, Bryant MP (1989) Syntrophomonas wolfei subsp. saponavida subsp. nov., a long chain fatty-acid degrading, anaerobic, syntrophic bacterium; Syntrophomonas wolfei subsp. wolfei subsp. nov.; and emended descriptions of the genus and species. Int J Syst Bacteriol 39:122–126CrossRefGoogle Scholar
  87. Ma AN (1999a) Treatment of palm oil mill effluent. Oil palm and environment: malaysia perspective. Malaysia Oil Palm Growers’Council, pp 277Google Scholar
  88. Ma AN (1999b) Innovations in management of palm oil mill effluent. The Planter Kuala Lumpur 75(881):381–389Google Scholar
  89. Ma AN (2000) Environmentalmanagement for the palm oil industry. PalmOil Dev 30:1–9Google Scholar
  90. Maia MRG, Chaudhary LC, Figueres L, Wallace RJ (2007) Metabolism of polyunsaturated fatty acids and their toxicity to the microflora of the rumen. Antonie Van Leeuwenhoek 91:303–314CrossRefGoogle Scholar
  91. Manish SR, Banerjee R (2008) Comparison of biohydrogen production processes. Int J Hydrogen Energy 33:279–286CrossRefGoogle Scholar
  92. McInerney MJ (1992) The genus Syntrophomonas, and other syntrophic bacteria. In: Balows A, Trüper HG, Dworkin M, Harder W, Schleifer KH (eds) The prokaryotes. Springer, New York, pp 2048–2057Google Scholar
  93. McInerney MJ, Bryant MP, Costerton JW (1981) Syntrophomonas wolfei gen. nov., an anaerobic, syntrophic, fatty acid-oxidizing bacterium. AppI Environ Microbiol 41:1029–1039Google Scholar
  94. McInery MJ, Bryant MP, Pfennig N (1979) Anaerobic bacterium that degrades fatty acids in syntrophic association with methanogens. Arch Microbiol 122:129–135CrossRefGoogle Scholar
  95. Mohammadi P, Ibrahim S, Annuar MSM, Ghafari S, Vikineswary S, Zinatizadeh AA (2011) Influences of environmental and operational factors on dark fermentive hydrogen production from wastes: a review. World Appl Sci J 13(2):188–199Google Scholar
  96. Morimoto M, Atsuko M, Atif AAY, Ngan MA, Fakhru’l-Razi A, Iyuke SE (2004) Biological production of hydrogen from glucose by natural anaerobic microflora. Int J Hydrogen Energy 29:709–713CrossRefGoogle Scholar
  97. Mumtaz T, Abd-Azizl S, Rahman NAA, Yee PL, Shirai Y, Hassan MA (2008) Pilot-scale recovery of low molecular weight organic acids from anaerobically treated palm oil mill effluent (POME) with energy integrated system. Afr J Biotechnol 7(21):3900–3905Google Scholar
  98. Najafpour GD, Zinatizadeh AAL, Mohamed AR, Isa MH, Nasrollahzadeh H (2006) High-rate anaerobic digestion of palm oil mill effluent in an upflow sludge-fixed film bioreactor. Process Biochem 41:370–379CrossRefGoogle Scholar
  99. Nath K, Das D (2004) Improvement of fermentative hydrogen production: various approaches. Appl Microbiol Biotechnol 65:520–529CrossRefGoogle Scholar
  100. Nielsen AT, Amandusson H, Bjorklund R, Dannetum H, Ejlertsson J, Ekedahl LG (2001) Hydrogen production from organic wastes. Int J Hydrogen Energy 26:547–550CrossRefGoogle Scholar
  101. Ntaikou G, Antonopoulou, Lyberatos G (2010) Biohydrogen production from biomass and wastes via dark fermentation: a review. Waste Biomass Valorization pp 1–19Google Scholar
  102. Nwuche CO, Ugoji EO (2008) Effects of heavy metal pollution on the soil microbial activity. Int J Environ Sci Technol 5(3):409–414Google Scholar
  103. Nwuche CO, Ugoji EO (2010) Effect of co-existing plant species on soil microbial activity under heavy metal stress. Int J Environ Sci Technol 7(4):697–704Google Scholar
  104. O-Thong S, Prasertsan P, Intrasungkha N, Dhamwichukorn S, Birkeland NK (2007) Improvement of biohydrogen production and treatment efficiency on palm oil mill effluent with nutrient supplementation at thermophilic condition using an anaerobic sequencing batch reactor. Enzyme Microb Technol 41:583–590CrossRefGoogle Scholar
  105. O-Thong S, Prasertsan P, Intrasungkha N, Dhamwichukorn S, Birkeland NK (2008a) Optimization of simultaneous thermophilic fermentative hydrogen production and COD reduction from palm oil mill effluent by Thermoanaerobacterium-rich sludge. Int J Hydrogen Energy 33:1221–1231CrossRefGoogle Scholar
  106. O-Thong S, Prasertsan P, Karakashev D, Angelidaki I (2008b) Thermophilic fermentative hydrogen production by the newly isolated Thermoanaerobacterium thermosaccharolyticum PSU-2. Int J Hydrogen Energy 33:1204–1214CrossRefGoogle Scholar
  107. Paillard D, McKain N, Chaudhary LC, Walker ND, Pizette F, Koppova I (2007) Relation between phylogenetic position, lipid metabolism and butyrate production by different Butyrivibrio-like bacteria from the rumen. Antonie Van Leeuwenhoek 91:417–422CrossRefGoogle Scholar
  108. Pavlostathis SG, Geraldogomez E (1991) Kinetics of anaerobic treatment. Water Sci Technol 24(8):35–59Google Scholar
  109. Pereira MA, Roest K, Stams AJM, Mota M, Alves MM, Akkermans A (2002) Molecular monitoring of microbial diversity in expanded granular sludge bed (EGSB) reactors treating oleic acid. FEMS Microbiol Ecol 41:95–103CrossRefGoogle Scholar
  110. Phang LY, Wakisaka M, Shirai Y, Hassan MA (2002) Freezing and thawing technique for the removal of suspended solids and concentration of palm oil mill effluent (POME). J Chem Eng Jpn 35(10):1017–1019CrossRefGoogle Scholar
  111. Phang LY, Hassan MA, Shirai Y, Wakisaka M, Abdul Karim MI (2003) Continuous production of organic acids from palm oil mill effluent with sludge recycle by the freezing-thawing method. J Chem Eng Jpn 36(6):707–710CrossRefGoogle Scholar
  112. Poh PE, Chong MF (2009) Development of anaerobic digestion methods for palm oil mill effluent (POME) treatment. Bioresour Technol 100:1–9CrossRefGoogle Scholar
  113. Rahim BA, Raj R (1982) Pilot plant study of a biological treatment system for palm oil mill effluent. In: Proceedings of regional workshop on palm oil mill technology and effluent treatment. PORIM, Malaysia, pp 163–170Google Scholar
  114. Redwood MD, Macaskie LE (2006) A two-stage, two-organism process for biohydrogen from glucose. Int J Hydrogen Energy 31(11):1514–1521CrossRefGoogle Scholar
  115. Rinzema A (1993) Anaerobic digestion of long chain fatty acids in UASB and expanded granular sludge bed reactors. Process Biochem 28:527–537CrossRefGoogle Scholar
  116. Roy F, Samain E, Dubourguier HC, Albagnac G (1986) Syntrophomonas sapovorans sp. nov., a new obligately proton reducing anaerobe oxidizing saturated and unsaturated long-chain fatty acids. Arch Microbiol 145:142–147CrossRefGoogle Scholar
  117. Rupani PF, Singh RP, Ibrahim MH, Esa N (2010) Review of current palm oil mill effluent (POME) treatment methods vermicomposting as a sustainable practice. World Appl Sci J 11(1):70–81Google Scholar
  118. Salehizadeh H, Van Loosdrecht MCM (2004) Production of polyhydroxyalkanoates by mixed culture: recent trends and biotechnical importance. Biotech Adv 22:261–279CrossRefGoogle Scholar
  119. Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61:262–280Google Scholar
  120. Shigematsu T, Tang Y, Mizuno Y, Kawaguchi H, Norimvra S, Kida K (2006) Microbial diversity of mesophilic methanogenic consortium that degrade longchain fatty acids in chemostat cultivation. J Biosci Bioeng 102:535–544CrossRefGoogle Scholar
  121. Singh RP, Embrandiri A, Ibrahim MH, Esa N (2011) Management of biomass residues generated from palm oil mill vermicomposting a sustainable option, resources. Conserv Recycling 55:423–434CrossRefGoogle Scholar
  122. Sousa DZ, Smidt H, Alves MM, Stams AJM (2007a) Syntrophomonas zehnderi sp. nov., an anaerobe that degrades long chain fatty acids in co-culture with Methanobacterium formicicum. Int J Syst Evol Microbiol 57:609–615CrossRefGoogle Scholar
  123. Sousa DZ, Pereira MA, Smidt H, Stams AJM, Alves MM (2007b) Molecular assessment of complex microbial communities degrading long chain fatty acids in methanogenic bioreactors. FEMS Microbiol Ecol 60:252–265CrossRefGoogle Scholar
  124. Sousa DZ, Pereira MA, Stams AJM, Alves MM, Smidt H (2007c) Microbial communities involved in anaerobic degradation of unsaturated or saturated long chain fatty acids. Appl Environ Microbiol 73:1054–1064CrossRefGoogle Scholar
  125. Sousa DZ, Alves JI, Alves MM, Smidt H, Stams AJM (2009) Effect of sulfate on methanogenic communities that degrade unsaturated and saturated long-chain fatty acids (LCFA). Environ Microbiol 11:68–80CrossRefGoogle Scholar
  126. Stronach SM, Rudd T, Lester JN (1978) Start-up of anaerobic bioreactors on high strength industrial wastes. Biomass 13:173–197CrossRefGoogle Scholar
  127. Stryer L (1995) Fatty acid metabolism, 4th edn. Biochemistry W.H. Freeman and Co., New York, pp 603–628Google Scholar
  128. Svetlitshnyi V, Rainey F, Wiegel J (1996) Thermosyntropha lipolytica gen. nov., sp. nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing short- and long-chain fatty acids in syntrophic coculture with a methanogenic archaeum. Int J Syst Bacteriol 46:1131–1137CrossRefGoogle Scholar
  129. Tabatabaei M, Zakaria MR, Rahim RA, Wright ADG, Shirai Y, Abdullah N, Sakai K, Ikeno S, Mori M, Kazunori M, Sulaiman A, Hassan MA (2009) Electron J Biotechnol 12(3):ISSN:0717-3458Google Scholar
  130. Tao YZ, Chen Y, Wu YQ, He YL, Zhou ZH (2007) High hydrogen yield from a two-step process of dark- and photo-fermentation of sucrose. Int J Hydrogen Energy 32(2):200–206CrossRefGoogle Scholar
  131. Thauer RK, Jungermann K, Decker K (1977) Energy conservation in chemotropic anaerobic bacteria. Bacteriol Rev 41:100–180Google Scholar
  132. Ting CH, Lee DJ (2007) Production of hydrogen and methane from wastewater sludge using anaerobic fermentation. Int J Hydrogen Energy 32:677–682CrossRefGoogle Scholar
  133. Tong SL, Jaafar AB (2006) POME Biogas capture, upgrading and utilization. Palm Oil Eng Bull 78:11–17Google Scholar
  134. Ueno Y, Tatara M, Fukui H, Makiuchi TT, Goto M, Sode K (2007) Production of hydrogen and methane from organic solid wastes by phase-separation of anaerobic process. Bioresour Technol 98:1861–1865CrossRefGoogle Scholar
  135. UNEP (1994) Treatment wastewater in the rubber industry. In: Cleaner production in the Asia-Pacific economic cooperation region. United Nations Environment Programme, Paris, pp 21–30Google Scholar
  136. Van der Leij FR, Witholt B (1995) Strategies for the suitable production of new biodegradable polyesters in plants: a review. Can J Microbiol 41(Suppl 1):222–238CrossRefGoogle Scholar
  137. Vijayaraghavan K, Ahmad D (2006) Biohydrogen generation from palm oil mill effluent using anaerobic contact filter. Int J Hydrogen Energy 31:1284–1291CrossRefGoogle Scholar
  138. Weiland P (2010) Biogas production current state and perspectives. Appl Microbiol Biotechnol 85:849–860CrossRefGoogle Scholar
  139. Weng C, Jeris JS (1976) Biochemical mechanisms in methane fermentation of glutamic and oleic acids. Water Res 10:9–18CrossRefGoogle Scholar
  140. Wong FM (1980) A review on the progress of compliance with the palm oil control regulations. Seminar on advances in palm oil effluent control technology, Kuala Lumpur, pp 142–149Google Scholar
  141. Wong PW, Nik MS, Nachiappa M, Balaraman V (2002) Pre-treatment and membrane ultra filtration using treated palm oil mill effluent (POME). Songklanakarin J Sci Tech 24(suppl):891–898Google Scholar
  142. Wong YS, Kadir MOAB, Teng TT (2009) Biological kinetics evaluation of anaerobic stabilization pond treatment of palm oil mill effluent. Bioresour Technol 100(21):4969–4975CrossRefGoogle Scholar
  143. Wu TY, Mohammad AW, Md Jahim J, Anuar N (2005) Penghasilan asid itakonik oleh Aspergillus terreus IMI 282743 dalam pelbagai jenis medium penghasilan. Pascasidang Kolokium Jabatan Kejuruteraan Kimia dan Proses. Malaysia, pp 147–151Google Scholar
  144. Wu C, Liu X, Dong X (2006) Syntrophomonas erecta subsp. Sporosyntropha subsp. nov., a spore-forming bacterium that degrades short chain fatty acids in co-culture with methanogens. Syst Appl Microbiol 29:457–462CrossRefGoogle Scholar
  145. Wu TY, Mohammad AW, Jahim JMd, Anuar N (2007) Palm oil mill effluent (POME) treatment and bioresources recovery using ultrafiltration membrane: effect of pressure on membrane fouling. Biochem Eng J 35:309–317CrossRefGoogle Scholar
  146. Wu TY, Mohammad AW, Jahim JM, Anuar N (2010) Pollution control technologies for the treatment of palm oil mill effluent (POME) through end-of-pipe processes. J Environ Manag 91:1467–1490CrossRefGoogle Scholar
  147. Lunar Engines (2009) Aerojet has completed assembly of a 5,500-pound-thrust liquid oxygen/liquid methane rocket engine: a propulsion technology under consideration as the way off the Moon for human explorers. Aviat Week Space Technol 171(2):16Google Scholar
  148. Yacob S, Hassan MA, Shirai Y, Wakisaka M, Subash S (2005) Baseline study of methane emission from open digesting tanks of palm oil mill effluent treatment. Chemosphere 59:1575–1581CrossRefGoogle Scholar
  149. Yacob S, Shirai Y, Hassan MA, Wakisaka M, Subash S (2006a) Startup operation of semi-commercial closed anaerobic digester for palm oil mill effluent treatment. Process Biochem 41:962–964CrossRefGoogle Scholar
  150. Yacob S, Shirai Y, Hassan MA, Wakisaka M, Subash S (2006b) Baseline study of methane emission from anaerobic pond of palm oil mill effluent treatment. Sci Total Environ 336:187–196Google Scholar
  151. Yee PL, Hassan MA, Shirai Y, Wakisaka M, Karim MIA (2003) Continuous production of organic acids from palm oil mill effluent with sludge recycle by the freezing–thawing method. J Chem Eng Jpn 36:707–710CrossRefGoogle Scholar
  152. Yokoi H (2002) Microbial production of hydrogen from starch manufacturing wastes. Biomass Bioenergy 22:389–395CrossRefGoogle Scholar
  153. Yu H (2002) Hydrogen production from rice winery wastewater in an upflow anaerobic reactor by using mixed anaerobic cultures. Int J Hydrogen Energy 27:1359–1365CrossRefGoogle Scholar
  154. Yusoff MZM, Hassan MA, Abd-Aziz S, Rahman NAA (2009) Start-Up of biohydrogen production from palm oil mill effluent under non-sterile condition in 50 L continuous stirred tank reactor. Int J Agric Res 4:163–168CrossRefGoogle Scholar
  155. Zakaria MR, Abd-Aziz S, Ariffin H, Rahman NAA, Yee PL, Hassan MA (2008) Comamonas sp. EB172 isolated from digester treating palm oil mill effluent as potential polyhydroxyalkanoate (PHA) producer. Afr J Biotechnol 7(22):4118–4121Google Scholar
  156. Zakaria MR, Tabatabaei M, Ghazali FM, Aziz SA, Shirai Y, Hassan MA (2010) Polyhydroxyalkanoates production from anaerobically trated palm oil mill effluent by new bacterial satrain Comamonas sp. EB172. World J Microbiol Biotechnol 26(5):767–774CrossRefGoogle Scholar
  157. Zhang T (2003) Biohydrogen production from starch in wastewater under thermophilic condition. J Environ Manage 69:149–156CrossRefGoogle Scholar
  158. Zhang CY, Liu XL, Dong XZ (2004) Syntrophomonas curvata sp. nov., an anaerobe that degrades fatty acids in co-culture with methanogens. Int J Syst Evol Microbiol 54:969–973CrossRefGoogle Scholar
  159. Zhao H, Yang D, Woese CR, Bryant MP (1993) Assignment of fattyacid-beta-oxidizing syntrophic bacteria to Syntrophomonadaceae fam. nov. on the basis of 16S rRNA sequence analysis. Int J Syst Bacteriol 43:278–286CrossRefGoogle Scholar
  160. Zinatizadeh AAL, Mohamed AR, Abdullah AZ, Mashitah MD, Husnain Isa M, Najafpour GD (2006) Process modeling and analysis of palm oil mill effluent treatment in an up-flow anaerobic sludge fixed film bioreactor using response surface methodology (RSM). Water Res 40:3193–3208CrossRefGoogle Scholar

Copyright information

© The Author(s) 2011

Authors and Affiliations

  • Anwar Ahmad
    • 1
  • Rumana Ghufran
    • 1
  • Zularisam Abd. Wahid
    • 1
  1. 1.Faculty of Civil Engineering and Earth ResourcesUniversity Malaysia Pahang (UMP)GambangMalaysia

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