INTRODUCTION

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Horizontal gene transfer in bacteria is an important factor in evolution. The interest in gene flux in the environment has also been stimulated by the discussion of the risks associated with the release of genetically modified microorganisms. In recent years, increased attempts have been made to assess the contribution of the various gene transfer mechanisms. Most effort has been put into plasmid-mediated conjugation and to a lesser extent into transformation, the uptake of free DNA. Phage-mediated transduction, however, has been widely neglected. One of the reasons may be that only a few generalized transducing phages from well-established laboratory strains are known.

In a previous study, we have shown that at least 95% of the natural strains of Salmonella enterica serovar Typhimurium harbor prophages (6). About 62% of the released phage strains could be assayed for their ability to transduce host genetic markers. With only two exceptions, all of them appeared to be generalized transducing phages. The detection of released phage particles from a lysogenic culture depends on the availability of suitable and sensitive bacterial indicator strains. The test for the transducing ability of the phages detected depends also on the availability of suitable mutant strains as recipients. Therefore, only 62% of the phages could be assayed.

Obviously, it would be rather unproductive to conduct a similar study for bacterial species that are not as well characterized and as well established in laboratories as serovar Typhimurium and its available mutant collections. If one asks for the overall occurrence of generalized transducing phages in natural habitats, still another problem interferes: 90 to 99.9% of all bacteria are considered unculturable (2). All these species would escape such an investigation. Therefore, it was desirable to develop a method for host-independent detection of transducing phage particles.

	MATERIALS AND METHODS

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Concentration of phage particles and destruction of free DNA. Lysates of phage P22H5 (c2 mutant) and of phage cI857 Sam7 were sterilized with membrane filters (pore size, 0.45 µm; Schleicher & Schuell, Dassel, Germany) and concentrated by ultracentrifugation (38.5-ml tubes; 2 h; 22,000 rpm; 5°C; Kontron TGA-50; rotor TST 28.38). The sediment was resuspended in 500 µl of DNase buffer (50 mM Tris-HCl [pH 7.5], 10 mM MgCl₂) and spiked with 1 µg of the control plasmid pHpaC (pUC19H with an amplifiable insert of phage ES18 DNA). To degrade nucleic acids, 100 µl of DNase I, 30 µl of RNase, and 30 µl of lysozyme were added (enzymes were from Boehringer, Mannheim, Germany) (stock solutions were at 10 mg ml¹) and shaken overnight at 6°C. The sample was extracted with an equal volume of chloroform and centrifuged for 30 min at 3,000 rpm at room temperature in a Heraeus Minifuge. The supernatant was incubated with 100 µl of lysozyme for 1 h at 37°C. After a further chloroform treatment and subsequent centrifugation, 100 µl of DNase, 30 µl of RNase, and 30 µl of lysozyme were added and incubated overnight at 6°C. Then, the phages were assumed to be ready for phenol extraction.

Environmental sample. From a sample of the supernatant of the mixed liquor in the aeration tank of an activated sludge treatment works (treatment plants Munich II, Munich, Germany) (samples are referred to as sewage water), coarse particles were removed by centrifugation (SS34, 5,000 rpm, 30 min; Sorvall) and filtration (Folded Filters; Schleicher & Schuell). Subsequently, the material was treated like phage lysates. Combined phage sediments of 1 liter of sewage water were resuspended in 3 ml of DNase buffer, and other volumes were adjusted accordingly.

Extraction of bacteriophage-encapsulated DNA. Phage-encapsulated DNA was extracted with the phenol-chloroform method (all components from Roth, Karlsruhe, Germany) according to the method of Maniatis et al. (4). DNA was precipitated at 80°C overnight by addition of 0.4 volumes of 5 M ammonium acetate and 2 volumes of 99% (vol/vol) ethanol. After centrifugation, DNA was washed with 70% (vol/vol) ethanol, dried under vacuum, and resolved overnight in double-distilled H₂O by shaking at 6°C. The DNA concentration was determined spectrophotometrically at 260 nm (Gene Quant; Pharmacia, Cambridge, United Kingdom). DNA quality and size were controlled electrophoretically.

PCR amplification. The amplification reaction mixture contained 10 ng of DNA, 5 µl of 10× PCR buffer, 1.5 mM MgCl₂ (for 16S ribosomal DNA [rDNA]) or 2.5 mM MgCl₂ (for plasmid control amplification), 2 U of native Taq polymerase (all components were from MBI Fermentas, St. Leon-Rot, Germany), 10 mM (each) deoxynucleotide triphosphates (PCR Nucleotide Mix; Boehringer) and 0.3 mM (each) primers. The final reactions/mixture volume was 50 µl. The mixture was incubated in a thermal cycler (RoboCycler; Stratagene). The cycling program was the following: initial denaturation at 95°C for 5 min; 30 cycles of 95°C for 1 min, primer annealing at 50°C for 1 min, and DNA synthesis at 72°C for 2 min; and a final extension step at 72°C for 7 min. All primers used in this study (Table 1) were synthesized by MWG Biotech (Ebersberg, Germany).

Cloning and sequencing. Amplification products were analyzed by electrophoresis in a 0.8% agarose gel, cloned into the plasmid pCR 2.1-TOPO by TA cloning (Invitrogen, Groningen, The Netherlands), and transformed into Escherichia coli strain TOP10F' (Invitrogen). Transformants were selected on Luria broth agar plates containing 100 µg of ampicillin ml¹, 50 µg of 5-bromo-4-chloro-3-indolyl--D-glucuronic acid ml¹, and 200 mM isopropyl--D-thiogalactopyranoside. Putative positive recombinant plasmids were isolated by boiling the plasmid preparations (3) and verified by EcoRI digestion (New England Biolabs, Beverly, Mass.). The plasmids selected for sequencing were purified using Jet Quick Clean-Up (Genomed, Bad Oyenhausen, Germany) following the manufacturer's recommendations. The plasmid DNA concentration was determined spectrophotometrically at 260 nm (Gene Quant; Pharmacia). Plasmid DNA (1.2 µg) was used for each sequencing reaction. Cloned 16S rDNAs were completely sequenced by M13 reverse, M13 (20) forward, and 525F primers (sequences of primers are shown in Table 1) using the DyeTerminator Cycle-Sequencing Kit AmpliTaq FS and the automatic sequence analyzer ABI PRISM 377 DNA Sequencer (Perkin-Elmer).

Nucleotide sequence accession numbers. Sequences have been submitted to GenBank. Accession numbers from AF324530 to AF324539 were assigned.

	RESULTS

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The concept. A lysate of a generalized transducing phage contains the entire genome of its propagating host strain as headful-sized fragments, thus representing a diluted complete gene library of the (killed) cells, including also the gene(s) for 16S rRNA. These 16S rRNA genes consist of highly conserved and variable sections, which are characteristic for the respective species or genus, like fingerprints. Therefore, we intended to extract phage-encapsulated DNA after removal of cells and complete destruction of all free nucleic acids, to amplify previously phage-encapsulated bacterial 16S rDNA by PCR. Primers used for amplification correspond with slight modifications to the primers fD1 and rP2, which were described elsewhere and allow amplification of almost full-length 16S rDNA of most eubacteria (7). After cloning of the amplification products, a collection of clones was expected to carry the 16S rDNA of those bacteria in the sample which have released general transducing phages. After the inserts are sequenced, the variable sections should yield information on which bacteria the transducing phages were released.

Removal of free DNA. The method was developed and conditions were optimized using a lysate of the generalized transducing Salmonella phage P22 as a test system.

View larger version (18K): [in this window] [in a new window]   FIG. 1.   Protocol for preparation and amplification of phage-encapsulated DNA.

View larger version (55K): [in this window] [in a new window]   FIG. 2.   Agarose gel analysis of PCR amplification products. We performed 16S rDNA amplification with primers 28F and 1492R and control insert amplification with standard primers M13 reverse and M13 (20) forward. Lane M, 1-kb ladder (New England Biolabs); lanes 1 and 5, amplification product of the control insert of plasmid pHpaC; lanes 2 to 4, test for the presence of the control insert of plasmid pHpaC or 16S rDNA, respectively, in phage lysates. All lysates were spiked with pHpaC, free DNA was degraded, and encapsulated DNA subsequently was extracted. Lane 2, control insert amplification of a P22 lysate; lane 3, 16S rDNA amplification of a P22 lysate; lane 4, 16S rDNA amplification of a lambda lysate. The 900-bp pHpaC amplification product is invisible at the applicated DNA amounts in lanes 1 and 5.

Application of the method to a sample of sewage water. Once the procedure was established with P22, we applied it to a sample of sewage water. To control reliability, spiking with the plasmid pHpaC was performed in parallel with P22 used as a positive control and phage lambda used as a negative control. Since both controls worked as expected and because no amplifiable pHpaC plasmid was left over after DNase treatment, all amplified material seen in an agarose gel was considered to derive from template DNA previously encapsulated in phage particles that were present in the sewage water.

	DISCUSSION

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The main problem in establishing a method for detection of transducible 16S rDNA in aquatic samples was the need for complete elimination of free bacterial DNA. This problem was solved by optimizing degradation conditions and by choosing a suitable Taq polymerase that was free of amplifiable 16S rDNA. Complete destruction of free DNA was confirmed by spiking the sample with a well-defined plasmid.

The amplified 16S rDNA from such samples was thought to derive from phage-encapsulated DNA. We cannot exclude the possibility that gene transfer agent-like elements, which have been found in Rhodopseudomonas capsulata, may also be involved (8). The process performed by these gene transfer agents resembles generalized transduction. But, if these particles are different from phages, they may be rare, and to our knowledge they have never been found in other species.

We have shown that this approach allows detection of generalized transducing phage particles without isolation and cultivation of the releasing host cells. Therefore, it will be possible also to detect transducing phages released from nonculturable species. Already the first application detected phages from four different gram-negative bacterial species. Half of the sequenced amplification products were most closely related to Aeromonas hydrophila, although for the genus Aeromonas, transducing phages have not yet been described. We emphasize, however, that the frequency of identified sequences does not necessarily reflect the quantitative composition of the bacterial community of the habitat. There may be dominant bacterial species which release no or only few transducing phages, while rarely represented species may use transduction as the main mechanism for gene transfer.

This method, which also worked well with samples from other habitats (unpublished data), will elucidate the natural hosts of generalized transducing phages in nature and the contribution of these phages to horizontal gene transfer among bacterial communities.

By application of suitable primers, this method could also detect viruses of higher organisms that are able to contribute to horizontal gene transfer among their hosts. It is known that retroviruses occasionally pick up cellular proto-oncogenes by excision errors. This process reflects the formation of specialized transducing particles of bacteriophage lambda and was detected by deleterious effects in the new host cells. Since retroviruses integrate themselves into the host genome at random positions, they may transfer other host genes in a similar way, which has not yet been detected, since they are less detrimental to their hosts. It is also possible that viruses other than retroviruses may perform some kind of general transduction as a consequence of erroneous DNA packaging.