A rapid DNA extraction method for PCR ... - Wiley Online Library

Letters in Applied Microbiology ISSN 0266-8254

ORIGINAL ARTICLE

A rapid DNA extraction method for PCR amplification from wetland soils J. Li1, B. Li1, Y. Zhou2, J. Xu2 and J. Zhao2 1 College of Life Sciences, Inner Mongolia University, Huhhot, China 2 College of Environment and Resources, Inner Mongolia University, Huhhot, China

Keywords ammonia-oxidizing archaea, ammoniaoxidizing bacteria, calcium chloride, glass bead, SDS method. Correspondence Ji Zhao, College of Environment & Resources, Inner Mongolia University, Huhhot 010021, China. E-mail: [email protected]

2010 ⁄ 1633: received 16 September 2010, revised 1 March 2011 and accepted 15 March 2011 doi:10.1111/j.1472-765X.2011.03047.x

Abstract Aims: We tested a method of rapid DNA extraction from wetland soil samples for use in the polymerase chain reaction. Methods and Results: The glass bead ⁄ calcium chloride ⁄ SDS method obtained in the present study was compared with the calcium chloride ⁄ SDS ⁄ enzymatic extraction method and the UltraClean Soil DNA Isolation Kit. Rapid DNA extraction could be completed within about two hours without purification steps. Conclusions: This study succeeded in establishing a fast soil DNA extraction protocol that can be applied to various environmental sources that are rich in humic acid content. Significance and Impact of the Study: The method provides a technology with high-quality DNA extraction from soils for testing the diversity of AOB and AOA.

Introduction Ammonia oxidation is the fist step in nitrification, a key process and limiting step in the global nitrogen cycle (Leininger et al. 2006). Ammonia-oxidizing bacteria (AOB) and the recently found ammonia-oxidizing organisms belonging to the archaeal domain (AOA) play important roles in the nitrogen cycle (You et al. 2009). Ammonia oxidation is now believed to be driven by these two major microbial groups. AOA have been found in various habitats including hot ⁄ thermal springs (Hatzenpichler et al. 2008), oceans (Beman et al. 2008), fresh water (Santoro et al. 2008) and soil (Tourna et al. 2008). However, they are very difficult to culture outside of these habitats because of their slow growth rates and their sensitivity to some organic substances. Up to now, only AOAs such as Nitrosopumilus maritimu, Nitrosocaldus yellowstonii, Nitrososphaera gargensis (You et al. 2009) were obtained. Cultivation-independent methods play important roles in helping us to understand the diversity and distribution of these microbes. Ammonia oxidation-related microbes are low in number and are hardly detectable using 16S rRNA (Junier et al. 626

2010). Therefore, alternative functional markers such as specific metabolic-pathway-related key enzymes, e.g., those involved in ammonia oxidation, have been used for ecological studies (Hermansson and Lindgren 2001). To amplify the amoA gene from AOA and AOB, an improved approach based on the dispersal of soils with glass beads is used to release ammonia-oxidizing microbes that are strongly adherent on soil colloids or located within the inner microporosity of soil aggregates (Robe et al. 2003). Many DNA extraction methods have been reported. SDS-based methods (Zhou et al. 1996) use CTAB or PVPP (Juniper et al. 1999) to remove humic substances; Al2(SO4)3 extraction methods (Dong et al. 2006; Persˇoh et al. 2008) use Al2(SO4)3 to remove humic substances and electroelution methods (Kallmeyer and Smith 2009) purify DNA directly extracted from marine sediments with an electroelution apparatus. DNA extraction solutions contain EDTA (Tsai and Olson 1992; Zhou et al. 1996; Miller et al. 1999; Martin-Laurent et al. 2001), which can combine with the divalent ions. It is very important to obtain nucleic acids from various environmental samples because DNA techniques allow less biased access to a greater portion of uncultivable microbes and

ª 2011 The Authors Letters in Applied Microbiology 52, 626–633 ª 2011 The Society for Applied Microbiology

J. Li et al.

also provide a useful tool for studying the structure and diversity of microbial communities (Robe et al. 2003). Many soil DNA extraction methods have been reported, such as a liquid nitrogen grinding method (Volossiouk et al. 1995), a microwave-based rupture method (Orsini and Romano-Spica 2001), an SDS-based method (Zhou et al. 1996), a bead-beating lysis method (Miller et al. 1999), a rapid freeze-and-thaw method (Tsai and Olson 1992), a cation-exchange method (Jacobsen and Rasmussen 1992), a solvent-based bead-beating method (Chen et al. 2006), an MS laboratory method (Martin-Laurent et al. 2001), a Nycodenz gradient separation method (He´le`ne et al. 2005), an Al2(SO4)3 extraction method (Persˇoh et al. 2008), our laboratory-devised calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method (Li et al. 2010) and our laboratory-devised glass bead ⁄ calcium chloride ⁄ SDS DNA extraction method, which was reported in this study. All DNA extraction methods focus on humic substances. To get high-purity DNA from soil, these methods generally include a subsequent purification step, such as Sepharose 4B, Sephadex G-200, Sephadex G-50 (Jackson et al. 1997) or electroelution (Kallmeyer and Smith 2009). In our laboratory-devised calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method, we also focused on these substances and obtained a method that is more efficient at removing humic acids from wetland soil because it uses a humic-substance-removal solution combined with calcium chloride solution. However, this method could only be used for 16S rDNA amplification and functional gene amplification from certain soils (in this study). The glass bead ⁄ calcium chloride ⁄ SDS DNA extraction method is based on the humic-substanceremoval technique derived from the calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method, and it can save time when used for functional gene amplification. In particular, ammonia oxidation-related microbes can be studied by this method. One commercial DNA purification kit and two laboratory-devised methods, a calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method (Li et al. 2010) and an improved glass bead ⁄ calcium ⁄ chloride ⁄ SDS method, were used to extract DNA directly from soil. The amoA gene and 16S rDNA were amplified to estimate the effectiveness of the different DNA extraction procedures. Materials and methods DNA extraction from soil The physicochemical properties of the four soils used in this study are presented in Table 1: Microbial biomass carbon according to the chloroform fumigation extraction method (Vance et al. 1987), organic carbon content in a Elementar Liqui TOC Analyzer (Germany), total nitrogen

A rapid DNA extraction method from wetland soils

Table 1 Properties of soil sample used in DNA extraction Soil properties

W1

W2

W3

W4

Size (lm) and % 1000 lm 500 lm 250 lm 100 lm 50 lm 10 lm 5 lm 2 lm Organic carbon (g kg)1) Total nitrogen (g kg)1) Microbial biomass carbon (mg kg)1)

100 99Æ89 96Æ8 68Æ62 47Æ23 16Æ62 7Æ86 1Æ298 70Æ69 2Æ115 987Æ01

100Æ00 99Æ67 90Æ31 53Æ37 33Æ12 13Æ70 6Æ49 0Æ93 62Æ60 2Æ037 731Æ21

100Æ00 100Æ00 91Æ56 30Æ71 15Æ28 3Æ79 0Æ98 0Æ00 25Æ08 1Æ089 246Æ35

100Æ00 100Æ00 96Æ41 53Æ14 30Æ05 6Æ67 2Æ22 0Æ00 2Æ37 1Æ7 756Æ10

content with semimicro-Kjeldahl determination (Cole et al. 1946), grain size distribution in a Microtrac S3500 (Montogomeryville, PA, USA). The experiment was conducted in the Inner Mongolian steppes (sites W1, W2) (4338¢N, 11642¢E) and Xilin River (sites W3, W4) (4408¢N, 11708¢E) in the Inner Mongolia Autonomous Region, China. Fresh soil samples were sieved (2 mm mesh) and stored at )20C. DNA was extracted from four soil samples using a commercial kit (UltraClean Soil DNA Isolation kit, Mobio Laboratories Inc., Carlsbad, CA, USA) according to the manufacturers’ recommendations (http://www.mobio. com ⁄ soil-dna-isolation ⁄ ultraclean-soil-dna-isolation-kit.html) and using two procedures developed in our laboratory. The steps of glass bead ⁄ calcium ⁄ chloride ⁄ SDS method and the calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method are as follows (see Figs 1 and 2, respectively). A humic-substance-removal solution containing 0Æ1 mol l)1 Tris, 0Æ1 mol l)1 Na4P2O7, 0Æ1 mol l)1 Na2EDTA, 1% PVP (w ⁄ v), 0Æ1 mol l)1 NaCl and 0Æ05% Triton X-100 (v ⁄ v), pH 10Æ0. DNA extraction buffer containing 0Æ1 mol l)1 Tris–HCl, 1Æ5 mol l)1 NaCl and 1% CTAB, pH 8Æ0. The mixture was homogenized using a VortexGenie 2 (Mobio Laboratories) for glass bead ⁄ calcium ⁄ chloride ⁄ SDS method. If the nucleic acid mix showed a white precipitate, 500 ll of a sterile, ice-cold carbonate dissolution mix (0Æ43 mol l)1 glacial acetic acid, 0Æ43 mol l)1 sodium acetate, and 0Æ17 mol l)1 sodium chloride, pH 4Æ6) (Kallmeyer and Smith 2009) was added, followed by incubation for 20 min on ice with 0Æ6 volume of isopropyl alcohol and centrifugation at 12 000 g for 5 min. Finally, 50 ll of TE buffer was added. PCR amplification of 16 S rDNA and the amoA gene To test the quality of the DNA extraction methods, 16S rDNA and amoA gene amplification was performed on


627


J. Li et al.

0·3 g soils

1 g soil + 2-ml centrifuge tube

2-ml centrifuge tube + three 2·5-mm-diameter glass beads or 3·0-mm-diameter glass beads

Homogenized and centrifuged at 12 000 g for 5 min at ambient temperature, the supernatant was then decanted

1 ml of a humic-substance-removal solution was added

1 ml of 0·5 mol l–1 CaCl2 solution was added, homogenized, and centrifuged at 12 000 g for 5 min

Homogenized for 2 min at maximum speed, centrifuged at 12 000 g for 2 min at ambient temperature, followed by decanting of the supernatant

The supernatant was then decanted, 1 ml of 0·05 mol l–1 sodium oxalate (pH 7·96) was added, homogenized and centrifuged at 12 000 g for 5 min and decanting of the supernatant 700 µl of DNA extraction buffer was added, then homogenized and 100 µl of 100 g l–1 lysozyme was added, mixed up and down several times and incubated for 30 min at 37°C 200 rpm in a table concentrator, then 200 µl 20% SDS was added and incubated for 1 h, centrifuged at 12 000 g

Calcium chloride solution was added (1 ml, 0·5 mol l–1), homogenized for 2 min at maximum speed, centrifuged at 12 000 g for 2 min at ambient temperature, followed by decanting of the supernatant

DNA extraction buffer (800 µl) was added 900-µl collected supernatant + equal volume of phenol-chloroform-isoamyl-alcohol (25 : 24 : 1) in the 2-ml centrifuge tube and centrifuged at 12 000 g

Homogenized for 5 s at maximum speed 200 µl 20% SDS was added, mixing up and down

800-µl collected supernatants + equal volume of phenol-chloroform-isoamyl-alcohol (25 : 24 : 1) in the 2-ml centrifuge tube and centrifuged at 12 000 g

and incubation for 10 min at 65°C, centrifuged at 12 000 g for 2 min 900-µl collected supernatant + equal volume of phenol-chloroform-isoamyl-alcohol (25 : 24 : 1) in the 2-ml centrifuge tube, centrifuged at 12 000 g for 5 min

700-µl collected supernatant + equal volume of chloroform-isoamyl-alcohol (24 : 1) in a 1·5-ml centrifuge tube and centrifuged at 12 000 g 600-µl collected supernatant + equal volume of chloroform-isoamyl-alcohol (24 : 1) in the 1·5-ml centrifuge tube and centrifuged at 12 000 g

800-µl collected supernatant + equal volume of chloroform-isoamyl-alcohol (24 : 1) in the 2-ml centrifuge tube, centrifuged at 12 000 g for 5 min

500-µl collected supernatant was incubated for 20 min on ice with 0·6 volume of isopropyl alcohol in the 1·5-ml centrifuge tube and centrifuged at 12 000 g

600-µl collected supernatant was incubated for 20 min on ice with 0·6 volume of isopropyl alcohol, centrifuged at 12 000 g for 5 min

The nucleic acids were washed with 70% ethanol and air-dried, followed by addition of 50 µl TE buffer

The nucleic acids were washed with 70% ethanol and air-dried, 50 µl of TE buffer was added Figure 1 The steps of glass bead ⁄ calcium ⁄ chloride ⁄ SDS method.

Figure 2 The steps of calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method.

DNA obtained directly from the four soils. Three replicates were analysed for the glass bead ⁄ calcium ⁄ chloride ⁄ SDS method. The 16S rDNA was amplified in a thermocycler from 1 ll of extracted soil DNA template with a total volume of 25 ll by using 2Æ0 ll of 2Æ5 mmol l)1 dNTP, 1Æ0 ll of 0Æ01 mol l)1 27F (5¢-AGA

GTT TGA TCM TGG CTC AG-3¢) (see Table 2), 1Æ0 ll of 0Æ01 mol l)1 1492R (5¢-TAC GGH TAC CTT GTT ACG ACT T-3¢) (see Table 2), 2Æ5 ll of 10· buffer (Promega, Madison, WI), and 0Æ2 ll of 5 U ll)1 Taq DNA polymerase under the following conditions: 5 min at 94C, 30 cycles of 30 s at 94C, 30 s at 55C, and 80 s at

628


J. Li et al.


Table 2 List of PCR primers Name

Sequence

References

Primer synthesis by

27F 1492R amoA-1F amoA-2R A189 A19F A643R

5¢-AGA GTT TGA TCM TGG CTC AG-3¢ 5¢-TAC GGH TAC CTT GTT ACG ACT T-3¢ 5¢-GGG GTT TCT ACT GGT GGT-3¢ 5¢-CCC CTC KGS AAA GCC TTC TTC-3¢ 5¢- GGN GAC TGG GAC TTC TGG-3¢ 5¢-ATG GTC TGG CTW AGA CG-3¢ 5¢- TCC CAC TTW GAC CAR GCG GCC ATC CA-3¢

Martin-Laurent et al. (2001) Martin-Laurent et al. (2001) Horz et al. (2004) Horz et al. (2004) Horz et al. (2004) Leininger et al. (2006) Treusch et al. (2005)

Invitrogen Invitrogen Invitrogen Invitrogen Invitrogen Invitrogen Invitrogen

72C, and an additional 10-min cycle at 72C. The 16S rDNA PCR products were then separated by electrophoresis on a 1% agarose gel. The AmoA gene of AOB was amplified using nested PCR. The first round of the nested PCR amplification from 2 ll of extracted soil DNA template was conducted in a total volume of 50 ll by using 4Æ0 ll of 2Æ5 · 10)3 mol l)1 dNTP, 1Æ0 ll of 1Æ0 · 10)9 mol l)1 A189 (5¢-GGN GAC TGG GAC TTC TGG-3¢) (see Table 2), 1Æ0 ll of 1Æ0 · 10)9 mol l)1 amoA-2R (5¢-CCC CTC KGS AAA GCC TTC TTC-3¢) (see Table 2), 5Æ0 ll of 10 · buffer (Promega) and 0Æ4 ll of 5 U ll)1 Taq under the following conditions: 5 min at 94C, 30 cycles of 40 s at 94C, 40 s at 55C, 40 s at 72C, and an additional 10-min cycle at 72C. The second round of the nested PCR amplification from 2 ll of extracted soil DNA template was conducted in a total volume of 50 ll using 4Æ0 ll of 2Æ5 · 10)3 mol l)1 dNTP, 1Æ0 ll of 1Æ0 · 10)9 mol l)1 amoA-1F (5¢-GGG GTT TCT ACT GGT GGT-3¢) (see Table 2) 1Æ0 ll of 1Æ0 · 10)9 mol l)1 amoA-2R (5¢-CCC CTC KGS AAA GCC TTC TTC-3¢) (see Table 2), 5Æ0 ll of 10 · buffer (Promega) and 0Æ4 ll of 5 U ll)1 Taq under the following conditions: 5 min at 94C, 30 cycles of 20 s at 94C, 20 s at 55C, 20 s at 72C, and an additional 10-min cycle at 72C. The AmoA gene of AOA was PCR amplified from 2 ll of extracted soil DNA template in a total volume of 50 ll using 8Æ0 ll of 2Æ5 · 10)3 mol l)1 dNTP, 1Æ0 ll of 1Æ0 · 10)9 mol l)1 A19F (5¢-ATG GTC TGG CTW AGA CG-3¢) (see Table 2), 1Æ0 ll of 1Æ0 · 10)9 mol l)1 A643R (5¢-TCC CAC TTW GAC CAR GCG GCC ATC CA-3¢) (see Table 2), 5Æ0 ll of 10 · buffer (Promega) and 0Æ4 ll of 5 U ll)1 Taq under the following conditions: 3 min at 95C, 35 cycles of 30 s at 94C, 30 s at 55C, 1 min at 72C, and an additional 10-min cycle at 72C. Cloning and sequencing PCR products of the AmoA gene from AOB were purified using a DNA purification kit (AxyPrep Biosciences, Hangzhou, China) according to the manufacturers’ recommendations, ligated into pMD-19T and then transformed into chemically competent Escherichia coli DH-5a (pro-

vided by the biochemistry laboratory of Inner Mongolia University, Huhhot, China). Clones were randomly selected for further analysis. Plasmid DNA was isolated from individual clones and sequenced by Invitrogen (Shanghai, China) using M13 forward and reverse primers. Phylogenetic analysis of AmoA Gene sequences from AOB were edited, and the vector sequences were deleted using the CLC Sequence Viewer 5 (http://www.clcbio.com). All sequences were analysed using megaBlast (http://blast.ncbi.nlm.nih.gov/Blast.cgi? PROGRAM=blastn&BLAST_PROGRAMS=megaBlast& PAGE_TYPE=BlastSearch&SHOW_DEFAULTS=on&LINK_ LOC=blasthome) to select the closest reference sequences, all amoA nucleotide sequences were aligned using Clustal X software, and an N-J tree (Jukes-Cantor correction) was constructed using mega software (Tamura et al. 2007). The bootstrap value was 1000 and the model was selected using the Kimura-2 parameter. AmoA gene sequence accession numbers The sequences identified in this study were submitted to GenBank under accession numbers HM481179–HM481197. Results Comparison of the three soil DNA extraction methods DNA was extracted from four soil samples using three methods (see the Materials and methods) (Fig. 3). The two methods devised in our laboratory do not contain EDTA in the DNA extraction buffer and use a humicsubstance-removal solution and calcium chloride solution to remove these substances before cell lysis. The glass bead ⁄ calcium chloride ⁄ SDS DNA extraction method obtains DNA that is about 23 kb in length, and the procedure can be completed within approximately two hours. In comparison with the calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method, which needs 4 h (Li et al. 2010), this method is faster. The glass bead ⁄ calcium


629


J. Li et al.

Ultra Clean Calcium Soil DNA Chloride-SDS Isolation -Enzymatic Kit Method

Glass Bead-Calcium Chloride-SDS method 4 4 4 3 3 3 2 2 2 1 1 1 M

4 3 2 1 4 3 2 1 23130 bp 9416 bp 6557 bp 4362 bp 2322 bp 2027 bp

564 bp

Figure 3 Gel electrophoresis of microbial genomic DNA (three duplicates) from four different soils by three methods.

chloride ⁄ SDS DNA extraction method saves time and can be used for PCR amplification (Fig. 4) such as that of 16S rDNA and especially for functional genes (amoA gene). Characterization of the new DNA extraction method The calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method (Li et al. 2010) was efficient in removing humic substances using the humic-substance-removal solution combined with the calcium chloride solution, which can combine with humic substances (Yuan et al. 2000; Davis

et al. 2002; Zhou et al. 2005). These processes are performed only in the first two steps (see the Materials and methods). As the subsequent steps include lysozyme treatment and heat treatment, they can lead to the release of humic substances from incompletely degraded animal and plant cells. For some soils, the humic substances released during the following steps under lysozyme and heating treatment cannot be removed. The sensitivity of PCR for DNA extracted from environmental samples is less than that for purified genomic DNA. This reduction could be attributed to humic substances or other interfering compounds present in the soil or sediments (Tsai and Olson 1992). Thus, the calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method could only be used to amplify 16S rDNA in all soils and the amoA gene from some of the soils in this study. Thus, the question remains: how to remove humic substances from incompletely degraded animal and plant cells? Larger glass beads were chosen to disperse the cells from the soil particles, allowing most dead animal and plant cells to release the humic substances into the humic-substance-removal solution and calcium chloride solution in the first two steps. Therefore, the humic substances are fully removed, and DNA obtained by the glass bead ⁄ calcium ⁄ chloride ⁄ SDS DNA extraction method can be used to amplify functional genes.

(a) 4

4 4

Round 2 3 3 3 2 2

2

1 1 1

4

4 4

3

Round 1 3 3 2 2

(b) 4 2

1

1 1

3

2

1

M

M 2000 bp 1000 bp 750 bp 500 bp 250 bp 100 bp

2000 bp 1000 bp 750 bp 500 bp 250 bp 100 bp

Figure 4 Gel electrophoresis of ammonia-oxidizing bacteria microbial genomic amoA and AOA microbial genomic gene fragment amplification from four different soils by glass bead ⁄ calcium chloride ⁄ SDS method.

Table 3 Results of PCR amplification of 16S rDNA, amoA gene fragments from ammoniaoxidizing bacteria (AOB) and ammoniaoxidizing archaea (AOA) carried out with the UltraClean Soil DNA Isolation Kit, calcium chloride ⁄ SDS ⁄ enzymatic and glass beads ⁄ calcium chloride ⁄ SDS method

Gene fragment 16S rDNA AOB amoA AOA amoA

Primer pair

Glass bead ⁄ calcium chloride ⁄ SDS method

Calcium chloride ⁄ SDS ⁄ enzymatic method

UltraClean soil DNA isolation kit

27F ⁄ 1492R A189 ⁄ 2Rw 1F ⁄ 2Rww A19F ⁄ A643R

++++ + ++++ ++++

++++ ) ++ )

++++ ) ) )

The single star represents round 1 of nested PCR, and the double star represents round 2 of nested PCR. ++++ means that PCR product of the target gene fragment can be obtained from four different soils by one of the three DNA extraction methods. +++ represents only three different soils, ++ only two difference soils, + only one soil, ) represents no PCR product of the target gene fragment can be obtained from four different soils by any one of the DNA extraction methods.

630


J. Li et al.


PCR amplification of 16S rDNA and the amoA gene The results of PCR amplification of 16S rDNA and amoA are presented in Table 3. The DNA obtained by the three DNA extraction methods from the four soils was used as a

template to amplify 16S rDNA (Fig. not shown). The specificity of the different primer combinations was tested, and the primer combination consisting of amoA-1F and amoA2R provided the most reliable performance in these studies (Rotthauwe et al. 1997). The AmoA gene was amplified

GU931351 clone-A-4|HM481182 72 clone-A-3|HM481181 clone-A-5|HM481183 99 clone-A-1|HM481179 63 clone-A-2|HM481180 61 GU225888 clone-C-6|HM481191 EU515194 Cluster 1 FJ853362 clone-B-2|HM481185 87 clone-B-3|HM481186 clone-B-1|HM481184 HM113507 66 82 GQ143593 87 clone-D-2|HM481194 66 97 GU931355 EU625025 GU931352 84 clone-B-5|HM481187 64 63 FJ890584 Nitrosospira-sp.LT2MFa|AY189145 clone-C-2|HM481188 100 65 clone-C-7|HM481192 100 FJ447346 Nitrosospira-multiformis|DQ228454 AF239880 52 100 clone-C-4|HM481190 99 clone-C-3|HM481189 Nitrosospira-sp.40KI|AJ298687 clone-D-1|HM481193 91 91 99 clone-D-4|HM481196 EU625184 clone-D-5|HM481197 65 clone-D-3|HM481195 100 AY249690 99 89 AB474979 Nitrosomonas-marina|AF272405 Nitrosomonas-oligotropha|AF272406 Nitrosomonas-ureae|AF272403 Nitrosomonas-eutropha|AJ298713 54 59

59 84

Cluster 2

Cluster 3

0·05 Figure 5 Phylogenetic tree based on amoA partial sequences was conducted using MEGA ver. 4Æ0 (Tamura, Dudley, Nei and Kumar 2007). Bootstrap values >50% are shown at each node. Numbers at each branch points indicate the percentage supported by bootstrap based on 1000 replicates. Bar indicates 0Æ05 substitution per nucleotide. ª 2011 The Authors Letters in Applied Microbiology 52, 626–633 ª 2011 The Society for Applied Microbiology

631


J. Li et al.

using nested PCR because of low abundance of AOB in our soil samples (Horz et al. 2004). Aliquots of the first round of PCR products were used as the templates for the second round of PCR (see the part of Materials and methods). The primer pairs selected were A189 ⁄ amo-2R and amo-1F ⁄ amoA-2R for the first and second rounds of nested PCR, respectively. The other primer pairs (Junier et al. 2010) were not tested in this study. The PCR product from the amoA gene of AOB was 491 bp. AmoA of AOB was amplified from only one of the four soil samples in the first round of nested PCR with the glass bead ⁄ calcium ⁄ chloride- ⁄ SDS DNA extraction method (Fig. 4a). However, the target bands were obtained from all four soil samples in the second round of nested PCR (Fig. 4a). AOB were not detected in soil samples W3 or W4 by the calcium chloride ⁄ SDS ⁄ enzyme DNA extraction method. The abundance and the composition of the indigenous bacterial community are dependent on the DNA recovery method used (Martin-Laurent et al. 2001), as clearly demonstrated by our results. The amoA gene of AOA could be amplified by general PCR from all soils using the primer pairs A19F ⁄ A643R (Treusch et al. 2005; Leininger et al. 2006) (Fig. 4b). This finding may indirectly show that the richness of AOA is higher than that of AOB. AmoA gene copies from Crenarchaeota (Archaea) are up to 3000-fold more abundant than bacterial amoA genes (Leininger et al. 2006). AOA amoA genes are more abundant, often as much as 80 times moreso than AOB amoA genes (Caffrey et al. 2007). Our results are consistent with the results of these previous studies. Phylogenetic relationships Four clone libraries of the amoA gene were constructed from the W1, W2, W3 and W4 soil samples. Five randomly selected clones were sequenced. A phylogenetic tree based on partial amoA sequences was constructed using mega ver. 4.0 (Tamura, Dudley, Nei, and Kumar 2007). In this study, 19 clones were obtained from the four soils: clones A-1 to A-5 belong to the W1 soil sample, clones B-1 to B-3 and B-5 belong to the W2 soil sample, clones C-2 to C-4, C-6 and C-7 belong to the W3 soil sample and clones D-1 to D-5 belong to the W4 soil sample. These clones received the accession numbers HM481179– HM481197 from GenBank, respectively (Fig. 5). The amoA sequences of AOB were divided into three clusters that were related to Nitrosospira. Nitrosomonas was not detected in these soil samples. Cluster 1, a Nitrosospira-like group, included five W1 clones, four W2 clones, one W3 clone and one W4 clone. Cluster 2 included two W3 clones. Cluster 3 included two W3 clones and four W4 clones. The AmoA gene sequences indicated that Nitrosospira of AOB may be the main nitrifiers in these four 632

different soils. Our results are consistent with the findings of Mohamed et al. (2010). Conclusion This study succeeded in establishing a fast soil DNA extraction protocol that can be applied to various environmental sources that are rich in humic acid content. In particular, the glass bead ⁄ calcium chloride ⁄ SDS method provides a rapid new approach for studying the diversity of AOB and ammonia-oxidizing archaea (AOA) in wetland and grassland soils. Acknowledgements This work was financially supported by the Key Project of National Programs for Fundamental Research and Development (2009CB125909). We thank Dr Alexander Buyantuyev for scientific paper writing classes, who is teaching in success Sino-US for Conservation, Energy and Sustainability Science. We thank our parents for the support and understanding these years. References Beman, J.M., Popp, B.N. and Francis, C.A. (2008) Molecular and biogeochemical evidence for ammonia oxidation by marine Crenarchaeota in the Gulf of California. ISME J 2, 429–441. Caffrey, J.M., Bano, N., Kalanetra, K. and Hollibaugh, J.T. (2007) Ammonia oxidation and ammonia-oxidizing bacteria and archaea from estuaries with differing histories of hypoxia. ISME J 1, 660–662. Chen, Y.C., Higgins, M.J., Maas, N.A. and Murthy, S.N. (2006) DNA extraction and Escherichia coli quantification of anaerobically digested biosolids using the competitive touchdown PCR method. Water Res 40, 3037–3044. Cole, J.E. and Pasks, C.R. (1946) Semi-micro Kjeldahl determination. Ann Chem 18, 61–65. Davis, C.J., Eschenazi, E. and Papadopoulos, K.D. (2002) Combined effects of Ca2+ and humic acid on colliod transport through porous media. Colloid Polym Sci 280, 52–58. Dong, D.X., Yan, A., Liu, H.M., Zhang, X.H. and Xu, Y.Q. (2006) Removal of humic substances from soil DNA using aluminium sulfate. J Microbiol Meth 66, 217–222. Hatzenpichler, R., Lebedeva, E.V., Spieck, E., Stoecker, K., Richter, A., Daims, H. and Wagner, M. (2008) A moderately thermophilic ammonia-oxidizing crenarchaeote from a hot spring. Proc Natl Acad Sci 105, 2134–2139. Helene, B., Franck, P., Van, T.V., Nathalie, L., Renaud, N., Timothy, M.V. and Pascal, S. (2005) High molecular weight DNA recovery from soils prerequisite for biotechnological metagenomic library construction. J Microbiol Methods 62, 1–11.


J. Li et al.

Hermansson, A. and Lindgren, P.E. (2001) Quantification of ammonia-oxidizing bacteria in arable soil by real-time PCR. Appl Environ Microbiol 67, 972–976. Horz, H.P., Barbrook, A., Field, C.B. and Bohannan, B.J.M. (2004) Ammonia- oxidizing bacteria respond to multifactorial global change. Proc Natl Acad Sci 101, 15136–15141. Jackson, C.R., Harper, J.P., Willoughby, D., Roden, E.E. and Churchill, P.F. (1997) A simple efficient method for the separation of humic substances and DNA from environmental samples. Appl Environ Microbiol 63, 4993–4995. Jacobsen, C.S. and Rasmussen, O.F. (1992) Development and application of a new method to extract bacterial DNA from soil based on separation of bacteria from soil with cationexchange resin. Appl Environ Microbiol 58, 2458–2462. Junier, P., Molina, V., Dorador, C., Hadas, O., Kim, O.S., Junier, T., Witzel, K.P. and Johannes, F.I. (2010) Phylogenetic and functional marker genes to study ammoniaoxidizing microorganisms (AOM) in the environment. Appl Microbiol Biotechnol 85, 425–440. Juniper, S.K., Cambon, M.A., Lesongeur, F. and Barbier, G. (1999) Extraction and purification of DNA from organic rich subsurface sediments. Mar Geol 174, 241–247. Kallmeyer, J. and Smith, D.C. (2009) An improved electroelutionmethod for separation of DNA from humic substances in marine sediment DNA extracts. FEMS Microbiol Ecol 69, 125–131. Leininger, S., Urich, T., Schloter, M., Schwark, L., Qi, J., Nicol, G.W., Prosser, J.I., Schuster, S.C. et al. (2006) Archaea predominate among ammonia- oxidizing prokaryotes in soils. Nature 442, 806–809. Li, J.Y., Zhao, J., Bian, Y., Wu, L.H. and Yu, J.L. (2010) DNA extraction and removing humic substance from wetland soil. Microbiol China 37, 1130–1137. Martin-Laurent, F., Philippot, L., Hallet, S., Chaussod, R., Germon, J.C., Soulas, G. and .Catroux, G. (2001) DNA extraction from soils: old bias for new microbial diversity analysis methods. Appl Environ Microbiol 67, 2354–2359. Miller, D.N., Bryant, J.E., Madsen, E.L. and Ghiorse, W.C. (1999) Evaluation and optimization of DNA extraction and purification procedures for soil and sediment samples. Appl Environ Microbiol 65, 4715–4724. Mohamed, N.M., Saito, K., Tal, Y. and Hill, R.T. (2010) Diversity of aerobic and anaerobic ammonia-oxidizing bacteria in marine sponges. ISME J 4, 38–48. Orsini, M. and Romano-Spica, V. (2001) A microwave-based method for nucleic acid isolation from environmental samples. Lett Appl Microbiol 33, 17–20. Persˇoh, D., Theuerl, S., Buscot, F. and Rambold, G. (2008) Towards a universally adaptable method for quantitative


extraction of high-purity nucleic acids from soil. J Microbiol Methods 75, 19–24. Robe, P., Nalin, R., Capellano, C., Vogel, T.M. and Simonet, P. (2003) Extraction of DNA from soil. Eur J Soil Biol 39, 183–190. Rotthauwe, J.H., Witzel, K.P. and Liesack, W. (1997) The ammonia monooxygenase structural gene amoA as a functional marker: molecular fine-scale analysis of natural ammonia-oxidizing populations. Appl Environ Microbiol 63, 4704–4712. Santoro, A.E., Francis, C.A., De-Sieyes, N.R. and Boehm, A.B. (2008) Shifts in the relative abundance of ammoniaoxidizing bacteria and archaea across physicochemical gradients in a subterranean estuary. Environ Microbiol 10, 1068–1079. Tamura, K., Dudley, J., Nei, M. and Kumar, S. (2007) MEGA4: Molecular Evolutionary Genetic Analysis (MEGA) software version 4.0. Mol Biol Evol 24, 1596–1599. Tourna, M., Freitag, T.E., Nicol, G.W. and Prosser, J.I. (2008) Growth activity and temperature responses of ammoniaoxidizing archaea and bacteria in soil microcosms. Environ Microbiol 10, 1357–1364. Treusch, A.H., Leininger, S., Kletzin, A., Schuster, S.C., Klenk, H.P. and Schleper, C. (2005) Novel genes for nitrite reductase and Amo-related proteins indicate a role of uncultivated mesophilic crenarchaeota in nitrogen cycling. Environ Microbiol 7, 1985–1995. Tsai, Y.L. and Olson, B.H. (1992) Detection of low numbers of bacterial cells in soils and sediments by polymerase chain reaction. Appl Environ Microbiol 58, 754–757. Vance, E.D., Brookes, P.C. and Jenkinson, D.S. (1987) An extraction method for measuring soil microbial biomass. Soil Biol Biochem 19, 703–707. Volossiouk, T., Robb, E.J. and Nazar, R.N. (1995) Direct DNA extraction for PCR- mediated assays of soil organisms. Appl Environ Microbiol 61, 3972–3976. You, J., Das, A., Dolan, E.M. and Hu, Z.Q. (2009) Ammoniaoxidizing archaea involved in nitrogen removal. Water Res 43, 1801–1809. Yuan, G., Theng, B.K.G., Parfitt, R.L. and Percival, H.J. (2000) Interactions of allophane with humic acid and cations. Eur J Soil Sci 51, 35–41. Zhou, J.Z., Bruns, M.A. and Tiedje, J.M. (1996) DNA recovery from soils of diverse composition. Appl Environ Microbiol 62, 316–322. Zhou, P., Yan, H. and Gu, B.H. (2005) Competitive complexation of metal ions with humic substances. Chemosphere 58, 1327–1337.


633