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We analyzed the temporal and spatial diversity of the microbiota in a low-usage and a high-usage hospital tap. We identified a tap-specific colonization pattern, with potential human pathogens being overrepresented in the low-usage tap. We propose that founder effects and local adaptation caused the tap-specific colonization patterns. Our conclusion is that tap-specific colonization represents a potential challenge for water safety.Humans are exposed to and consume large amounts of tap water in their everyday life, with the tap water microbiota representing a potent reservoir for pathogens (8). Despite the potential impact, our knowledge about the ecological diversification processes of the tap water microbiota is limited (4, 11).The aim of the present work was to determine the temporal and spatial distribution patterns of the planktonic tap water microbiota. We compared the summer and winter microbiota from two hospital taps supplied from the same water source. We analyzed 16S rRNA gene clone libraries by using a novel alignment-independent approach for operational taxonomic unit (OTU) designation (6), while established OTU diversity and richness estimators were used for the ecological interpretations.Tap water samples (1 liter) from a high-usage kitchen and a low-usage toilet cold-water tap in Akershus University Hospital, Lørenskog, Norway, were collected in January and July 2006. The total DNA was isolated and the 16S rRNA gene PCR amplified and sequenced. Based on the sequences, we estimated the species richness and diversity, we calculated the distances between the communities, and trees were constructed to reflect the relatedness of the microbiota in the samples analyzed. Details about these analytical approaches are given in the materials and methods section in the supplemental material.Our initial analysis of species composition was done using the RDPII hierarchical classifier. We found that the majority of pathogen-related bacteria in our data set belonged to the class Gammaproteobacteria. The genera encompassed Legionella, Pseudomonas, and Vibrio (Table (Table1).1). We found a significant overrepresentation of pathogen-related bacteria in the toilet tap (P = 0.04), while there were no significant differences between summer and winter samples. Legionella showed the highest relative abundance for the pathogen-related bacteria. With respect to the total diversity, we found that Proteobacteria dominated the tap water microbiota (representing 86% of the taxa) (see Table S1 in the supplemental material). There was, however, a large portion (56%) of the taxa that could not be assigned to the genus level using this classifier.
TABLE 1.
Cloned sequences related to human pathogensaOpen in a separate windowaThe relatedness between the cloned sequences and potential pathogens was determined by BLAST searches of the NCBI database, carried out using default settings.To obtain a better resolution of the uncharacterized microbiota, we analyzed the data using a clustering approach that is not dependent on a predefined bacterial group (see the materials and methods section in the supplemental material for details). These analyses showed that there were three relatively tightly clustered groups in our data set (Fig. (Fig.1A).1A). The largest group (n = 590) was only distantly related to characterized betaproteobacteria within the order Rhodocyclales. We also identified another large betaproteocaterial group (n = 320) related to Polynucleobacter. Finally, a tight group (n = 145) related to the alphaproteobacterium Sphingomonas was identified.Open in a separate windowFIG. 1.Tap water microbiota diversity, determined by use of a principal component analysis coordinate system. (A) Each bacterium is classified by coordinates, with the following color code: brown squares, kitchen summer; red diamonds, toilet summer; green triangles, kitchen winter; and green circles, toilet winter. (B and C) Each square represents a 1 × 1 (B) or 5 × 5 (C) OTU. PC1, first principal component; PC2, second principal component.The tap-specific distributions of the bacterial groups were investigated using density distribution analyses. A dominant population related to Polynucleobacter was identified for the toilet summer samples, while for the winter samples there was a dominance of the Rhodocyclales-related bacteria. The kitchen summer samples revealed a dominance of Sphingomonas. The corresponding winter samples did not reveal distinct high-density bacterial populations (see Table S2 in the supplemental material).Hierarchical clustering for the 1 × 1 OTU density distribution confirmed the relatively low overlap for the microbiota in the samples analyzed (Fig. (Fig.2).2). We found that the microbiota clustered according to tap and not season.Open in a separate windowFIG. 2.Hierarchical clustering for the density distribution of the tap water microbiota. The density of 1 × 1 OTUs was used as a pseudospecies for hierarchical clustering. The tree for the Cord distance matrix is presented, while the distances calculated using the three distance matrices Cord, Brad Curtis, and Sneath Sokal, respectively, are shown for each branch.We have described the species diversity and richness of the microbiota in Table S3 in the supplemental material. For the low taxonomic level, these analyses showed that the diversity and species richness were greater for the winter samples than for the summer samples. Comparing the two taps, the diversity and richness were greater in the kitchen tap than in the toilet tap. In particular, the winter sample from the kitchen showed great richness and diversity. The high taxonomic level, however, did not reveal the same clear differences as did the low level, and the distributions were more even. Rarefaction analyses for the low taxonomic level confirmed the richness and diversity estimates (see Fig. S1 in the supplemental material).Our final analyses sought to fit the species rank distributions to common rank abundance curves. Generally, the rank abundance curves were best fitted to log series or truncated log normal distributions (see Table S4 in the supplemental material). The log series distribution could be fit to all of the samples except the kitchen summer samples at the low taxonomic level, while the truncated log normal distribution could not be fit to the kitchen samples at the high taxonomic level. Interestingly, however, the kitchen winter sample was best fit to a geometric curve at both the high and the low taxonomic level.Diversifying, adaptive biofilm barriers have been documented for tap water bacteria (7), and it is known that planktonic bacteria can interact with biofilms in an adaptive manner (3). On the other hand, tap usage leads to water flowthrough and replacement of the global with the local water population by stochastic founder effects (1).Therefore, we propose that parts of the local diversity observed can be explained by local adaptation (10) and parts by founder effects (9).Most prokaryote diversity measures assume log normal or log series OTU dominance density distributions (5). The kitchen winter sample, however, showed deviations from these patterns by being correlated to geometric distributions (in addition to the log series and truncated log normal distributions for the high taxonomic level). This sample also showed a much greater species richness than the other samples. A possible explanation is that the species richness of the tap water microbiota can be linked to usage and that the kitchen tap is driven toward a founder microbiota by high usage.Since our work indicates an overrepresentation of Legionella in the low-usage tap, it would be of high interest to determine whether the processes for local Legionella colonization can be related to tap usage. Understanding the ecological forces affecting Legionella and other pathogens are of great importance for human health. At the Akerhus University Hospital, this was exemplified by a Pseudomonas aeruginosa outbreak in an intensive care unit, where the outbreak could be traced back to a single tap (2). 相似文献6.
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Paul W. J. J. van der Wielen Gertjan Medema 《Applied and environmental microbiology》2010,76(14):4876-4881
Bacteroidales species were detected in (tap) water samples from treatment plants with three different PCR assays. 16S rRNA gene sequence analysis indicated that the sequences had an environmental rather than fecal origin. We conclude that assays for Bacteroidales 16S rRNA genes are not specific enough to discern fecal contamination of drinking water in the Netherlands.Drinking water in many countries is routinely monitored for recent fecal contamination by testing for fecal indicator organisms Escherichia coli, thermotolerant coliforms, and/or intestinal enterococci to demonstrate microbial safety (13, 21, 42). Although these indicator organisms have been used for many decades, they have some limitations: the number of E. coli/coliform/enterococcus bacteria in feces is relatively low (18, 38), and they sometimes might be able to grow in the environment (10, 11, 14, 27). Consequently, scientists have been searching for alternative indicator organisms to determine fecal contamination of water. In 1967, bacteria belonging to the genus Bacteroides were suggested as alternative indicator organisms (26). Bacteroides spp. might have some advantages over the traditional indicator organisms. The numbers of Bacteroides spp. in the intestinal tract of humans and animals are 10 to 100 times higher than the numbers of E. coli or intestinal enterococci (1, 2, 12, 26). However, the use of Bacteroides spp. as indicator organisms was hampered by the complex cultivation conditions required (1, 2). The introduction of molecular methods made it possible to detect bacterial species that belong to the order Bacteroidales, an order that includes the genus Bacteroides, without cultivation. As a result, real-time PCR methods were developed for the quantitative detection of Bacteroidales in surface and recreation water and the potential of Bacteroidales species as an indication of fecal contamination of recreational waters was demonstrated (6, 12, 16, 19, 20, 29). Bacteroidales species might be useful indicator organisms for fecal contamination of drinking water as well. However, methods to detect fecal contamination in drinking water should be more sensitive, because people ingest more drinking water and the quality assessments and standards for fecal contamination are stricter than for bathing water. Studies exploring real-time PCR for the detection of Bacteroidales genes in drinking water have not been published to our knowledge. The objective of our study was, therefore, to determine if assays for the detection of Bacteroidales 16S rRNA genes can be used to detect fecal contamination in drinking water.Unchlorinated tap water samples were obtained in November 2007 and February 2010 from one or more locations in the distribution systems of nine different drinking water treatment plants (plants A to I; Table Table1)1) that produced unchlorinated drinking water from confined (plants B, C, E, F, and G) and unconfined (plants A, D, H, and I) groundwater. The treatment plants are located in the central part of the Netherlands within 90 km of each other. In addition, untreated groundwater from extraction wells and/or untreated raw groundwater (mixture of groundwater from different extraction wells) was sampled in March 2008 (Table (Table1).1). Water samples (100 ml) were filtered over a 25-mm polycarbonate filter (0.22-μm pore size, type GTTP; Millipore, Netherlands) and a DNA fragment was added as internal control to determine the recovery efficiency of DNA isolation and PCR analysis (2a, 40). DNA was isolated using a FastDNA spin kit for soil (Qbiogene, United States) according to the supplier''s protocol. Primer sets AllBac 296f and AllBac 412r, resulting in a PCR product of 108 bp, were used in combination with TaqMan probe AllBac375Bhqr to quantitatively determine the number of Bacteroidales 16S rRNA gene copies in the water samples using a real-time PCR instrument (20). The PCR cycle after which the fluorescence signal of the amplified DNA was detected (threshold cycle [CT]) was used to quantify the concentration of 16S rRNA gene copies. Quantification was based on comparison of the sample CT value with the CT values of a calibration curve graphed using known copy numbers of the Bacteroidales 16S rRNA gene, as previously described (12, 20). The correlation coefficient of the calibration curve was 0.99, and the efficiency of the PCR 95 to 105%. Finally, the Bacteroidales cell number was calculated by using the recovery rate of the internal standard and assuming five 16S rRNA gene copy numbers per cell (5). The detection limit of this gene assay was 50 Bacteroidales cells 100 ml−1 (corresponding to 10 16S rRNA gene copies per reaction mixture). Furthermore, the 16S rRNA genes that were obtained from several water samples from treatment plant C with the AllBac and TotBac (12) primer sets were sequenced, and the nearest relatives were obtained from the GenBank database using BLAST searches.
Open in a separate windowaValues are the average results and standard deviations from replicate PCRs on the same water sample using the AllBac primer set (20). In November 2007, the distribution systems (tap water) of plants A, B, and G were sampled at three different locations, whereas for the other plants, one location in the distribution system was sampled. In March 2008, raw water of plants A to G was sampled, as well as one (plant E) or three (plant C) different extraction wells. Finally, in February 2010, the distribution systems of plants A, B, C, D, E, and G were sampled again.bMore than one tap water sample from a treatment plant means that samples were taken at different locations in the distribution system.The Bacteroidales 16S rRNA gene, quantified with the AllBac primer set, was detected in all tap water samples in November 2007 and February 2010. The number of cells varied between 154 and 7,862 Bacteroidales cells 100 ml−1, and the numbers in tap water of each plant were similar in 2007 and 2010 (Table (Table1).1). The Bacteroidales counts were high compared to the number of E. coli that are occasionally observed in fecally contaminated drinking water (17a) but low compared to numbers observed in surface water (4, 20, 22). Water from the extraction wells and raw water used for unchlorinated drinking water production were analyzed, and Bacteroidales species were detected in 10 out of 15 samples (Table (Table1).1). These results would imply that the extracted groundwater, raw water, and tap water were fecally contaminated. According to the Dutch drinking water decree (2b), both raw and tap water from the nine different treatment plants are regularly analyzed for fecal contamination by monitoring for E. coli, F-specific RNA phages, and somatic coliphages. For at least the last 10 years, these indicator organisms have not been detected in these waters.Additional qualitative PCR analyses using TotBac and BacUni primer sets (12, 19) targeting other parts of the Bacteroidales 16S rRNA gene were performed to confirm the presence of Bacteroidales species in the water samples of November 2007 and March 2008. Nine or 10 of the 11 samples that were positive with the AllBac primer set were also positive with the TotBac and BacUni primer sets (data not shown). The BacUni primer set has a higher detection limit (30 gene copies per PCR; 19), which could explain the difference from the results with the AllBac primer set. The TotBac primer set has the same detection limit as the AllBac primer set (12), but small differences in PCR efficiencies might have resulted in different results, since some water samples showed Bacteroidales 16S rRNA gene copy numbers around the detection limit (Table (Table1).1). Nevertheless, the additional PCR analyses demonstrated that the detection of Bacteroidales species in tap, raw, and extracted well water with the AllBac primer set was not an artifact. The primer sets used were developed in three different studies (12, 19, 20) but have been applied in a number of recent studies to detect fecal contamination of surface water (3, 4, 16, 22, 33, 34). The results from most of these studies showed that 16S rRNA genes of Bacteroidales were present in all surface water samples tested. Only Sinigalliano et al. (34) observed that 2 out of 4 water samples were negative with the TotBac primer set. However, the detection limit of the assay was not specified in that study.The nine different treatment plants tested in our study produce unchlorinated drinking water from groundwater, which is considered to be of high hygienic quality. In addition, the extraction wells are protected from fecal contamination by a protection zone where no activities related to human waste or animal manure are allowed. In the Netherlands, this protection zone is based on a 60-day residence time of the water. Previous studies have demonstrated that a residence time of 60 days is highly effective in removing fecal bacteria and viruses (30, 31, 39). Moreover, the Bacteroidales numbers in tap water in November 2007 were significantly higher than the numbers in raw groundwater in March 2008 (Mann-Whitney U test; P < 0.01). Because the recovery efficiency of the internal control was the same between raw water and tap water samples, this result demonstrates that Bacteroidales cell numbers increased during treatment and/or drinking water distribution. This result could suggest that the water was fecally contaminated during drinking water treatment and/or distribution. However, it is unlikely that the integrity of nine different treatment trains and/or supply systems was affected in the sampling period. The statutory monitoring did not show the presence of E. coli at these sites. Another hypothesis is that the increase of Bacteroidales cell numbers in tap water was caused by the growth of Bacteroidales species in (drinking) water systems. In summary, it is unexpected that the majority of the tap water, raw water, and extracted groundwater samples were fecally contaminated. These unexpected observations raise the question of whether the PCR methods detect only fecal Bacteroidales species and, thus, if the gene assays are suitable to discern fecal contamination in drinking water in the Netherlands.Sequence analyses of the Bacteroidales 16S rRNA genes were performed to determine the relatedness of sequences from the different sampling sites to sequences from the nearest relatives in the GenBank database. All sequences contained the primer regions, indicating that nonspecific amplification had not occurred in the PCRs. Because the PCR product from the AllBac primer set was small (108 bp), many 16S rRNA gene sequences (100 to 5,000) in the GenBank database were identical to the Bacteroidales 16S rRNA gene sequences obtained from groundwater and unchlorinated tap water samples from plant C. These identical 16S rRNA gene sequences were in general obtained from fecal sources, but some of them came from environmental rather than fecal sources (Table (Table2).2). The AllBac 16S rRNA gene sequences from tap water and groundwater had relative high similarities (96.3 to 100%) to sequences from bacterial species of the genera Bacteroides, Prevotella, and Tannerella (Table (Table2),2), which all belong to the order Bacteroidales.
TABLE 1.
Numbers of Bacteroidales cells in extraction wells, raw groundwater, and unchlorinated tap water of nine different groundwater plants in the NetherlandsaPlant | Source of sample | No. (100 ml−1) of Bacteroidales cells in: | ||
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2007 | 2008 | 2010 | ||
A | Tap water 1b | 5,948 ± 950 | ||
Tap water 2 | 2,682 ± 1,459 | 1,254 ± 216 | ||
Tap water 3 | 4,362 ± 947 | 439 ± 136 | ||
Raw water | 96 ± 15 | |||
B | Tap water 1 | 3,553 ± 981 | 5,302 ± 2,952 | |
Tap water 2 | 4,487 ± 391 | 2,119 ± 1,367 | ||
Tap water 3 | 7,862 ± 4,588 | 3,896 ± 3,003 | ||
Raw water | 3,209 ± 833 | |||
C | Tap water 1 | 661 ± 75 | 386 ± 199 | |
Tap water 2 | 1,051 ± 626 | |||
Tap water 3 | 831 ± 584 | |||
Tap water 4 | 1,254 ± 216 | |||
Extraction well 1 | 1,126 ± 262 | |||
Extraction well 2 | 2,666 ± 51 | |||
Extraction well 3 | <50 | |||
Raw water | 90 ± 44 | |||
D | Tap water | 1,103 ± 29 | 1,254 ± 216 | |
Raw water | 48 ± 16 | |||
E | Tap water | 1,302 ± 222 | 1,254 ± 216 | |
Extraction well 1 | 671 ± 97 | |||
F | Tap water | 1,317 ± 198 | ||
Raw water | <50 | |||
G | Tap water 1 | 675 ± 92 | 439 ± 300 | |
Tap water 2 | 216 ± 65 | 249 ± 98 | ||
Tap water 3 | 154 ± 6 | 322 ± 137 | ||
Raw water | <50 | |||
H | Tap water | 7,073 ± 845 | ||
Raw water | 511 ± 254 | |||
I | Tap water | 1,577 ± 176 | ||
Raw water | 420 ± 66 |
TABLE 2.
Nearest relatives in GenBank to the Bacteroidales 16S rRNA gene sequences obtained from groundwater and unchlorinated tap water from plant C using different primer setsaOpen in a separate windowaPrimer sets AllBac (20) and TotBac (12) were used in PCRs of samples, and GenBank was searched for relatives using BLAST.bOTUs are indicated by the values in parentheses (number of sequences belonging to the OTU/total number of sequences analyzed).cNumber of base pairs identical in both sequences/total number of base pairs in sequences.16S rRNA gene sequences obtained with the TotBac primer set were longer (∼370 bp) and did not show 100% similarity with the nearest relatives in the GenBank database (Table (Table2).2). Sequences from the GenBank database that showed the highest similarity (91.6% to 99.7%) with the 16S rRNA gene sequences from tap water and groundwater from plant C were in general isolated from environmental sources (Table (Table2).2). The 16S rRNA gene sequences from cultivated bacterial species that showed the highest similarity to the 16S rRNA gene sequences obtained in our study belonged to different genera (Table (Table2).2). Some of these genera (Salinimicrobium, Xanthobacillum, and Psychroserpens) did not belong to the order Bacteroidales. However, the 16S rRNA gene sequences from bacterial species of these genera showed low similarities with the sequences obtained in this study (83.2% to 90.1%) and six mismatches to the TotBac primers. Thus, it is unlikely that DNA from bacterial species belonging to Salinimicrobium, Xanthobacillum, and Psychroserpens was amplified in the gene assay. More importantly, the majority of the nearest environmental clone sequences retrieved from the GenBank database showed no or a single mismatch with the AllBac and TotBac primer and probe sequences. Thus, these primer sets are capable of amplifying 16S rRNA genes from bacteria that have been observed in ecosystems outside the intestinal tract of humans and animals.16S rRNA gene sequences related to Prevotella species were commonly observed in extracted groundwater, raw water, and tap water (Table (Table2).2). The isolation of Prevotella paludivivens from rice roots in a rice field soil (35) demonstrated the environmental nature of some Prevotella species. In addition, primer sequences developed for the detection of fecal Bacteroidales species (8, 12, 19, 20, 25, 29) showed no or a single mismatch with 16S rRNA gene sequences from P. paludivivens, Xylanibacterium oryzae, Paludibacter propionicigenes, Proteiniphilum acetatigenes, and Petrimonas sulfuriphila that are present in the GenBank database. These five Bacteroidales species have all been isolated from ecosystems other than the gastrointestinal tract. Consequently, primer sets for 16S rRNA genes of Bacteroidales species cannot always be used to discern fecal contamination in water.A number of 16S rRNA gene sequences observed in groundwater and tap water fell in the genus Bacteroides. The presence of Bacteroides 16S rRNA gene sequences in groundwater and tap water might also suggest that some Bacteroides species are capable of growth in the environment. However, until now, type strains of Bacteroides species growing outside the animal intestinal tract have not been published. Another possible explanation is that the observed 16S rRNA gene sequences originate from Bacteroides species that inhabit the anoxic intestinal tract of insects. Previous studies have shown that bacterial species belonging to the genus Bacteroides are common inhabitants of the hindguts of insects (15, 23, 24, 28, 32). Some of the 16S rRNA gene sequences obtained with the AllBac primer set in our study showed 100% similarity to 16S rRNA gene sequences from the hindgut of insects. Moreover, a number of 16S rRNA gene sequences isolated from the hindguts of insects (15, 23, 24, 32) showed no or a single mismatch with the TotBac and AllBac primer and probe sequences. In conclusion, these primer sets are capable of detecting Bacteroides species from the hindgut of insects as well. Water insects are normal inhabitants of groundwater and drinking water distribution systems (7, 41) and might be a source of Bacteroides species in water. Bacteroides species from insect feces do not indicate fecal pollution by warm-blooded animals, and insects do not normally shed human fecal pathogenic microorganisms. Bacteroides species from insect feces, therefore, can hamper Bacteroides gene assays developed for the detection of water fecally contaminated by warm-blooded animals. Additional cultivation techniques in combination with molecular tools are required to demonstrate the persistence or growth of Bacteroides bacteria in groundwater and drinking water or whether Bacteroides bacteria are present in water insects. However, these experiments were beyond the scope of our study.The three extraction wells of plant C are located close to each other and extract water from the same aquifer. Subsequently, extracted water from the three wells is mixed and enters the treatment plant as raw water. We hypothesize that if a fecal source in the vicinity of the extraction field of plant C contaminated the groundwater, water from the extraction wells and raw water should (partly) have the same Bacteroidales species. Although a relatively limited amount of clones was sequenced per sample (16), the diversity of Bacteroidales operational taxonomic units (OTU) within a sample was low (Table (Table2).2). In contrast, unique 16S rRNA gene sequences were observed between the different water types (e.g., extracted groundwater, raw water, and tap water) and sequence overlap between water types was low. These results demonstrate that the Bacteroidales 16S rRNA gene sequences at the sampling locations were not from the same fecal source and imply once again that Bacteroidales species were environmental rather than fecal.Finally, we hypothesized that if the Bacteroidales species observed in tap water were of nonfecal origin, human- and/or bovine-specific Bacteroidales strains should not be present in tap water. We tested for the presence of human- or bovine-specific Bacteroidales strains by using source-specific 16S rRNA gene assays (5) on tap water samples from February 2010. The results showed that human- and bovine-specific Bacteroidales 16S rRNA genes could not be detected in tap water, whereas a PCR product was always detected with the positive control. Again, these results indicate that the Bacteroidales species observed in tap water were of nonfecal origin.Overall, the results from our study indicate that gene assays for Bacteroidales detected environmental rather than fecal Bacteroidales species in groundwater and tap water from treatment plants in the Netherlands. First, Bacteroidales 16S rRNA gene sequences obtained from water samples taken at plant C showed (high) similarity to clone sequences that were isolated from environmental sources. The majority of these clone sequences and several Bacteroides clone sequences from the hindguts of insects showed no or a single mismatch with AllBac, TotBac, and BacUni primer and probe sequences. Second, the primer and probe sequences used for the gene assays have no or a single mismatch with 16S rRNA gene sequences of environmental Bacteroidales species P. paludivivens, X. oryzae, P. propionicigenes, P. acetatigenes, and/or P. sulfuriphila (9, 17, 35-37). Third, Bacteroidales 16S rRNA gene sequences from raw water and water from extraction wells were unique, and sequence overlap between water types was low. It is expected that in the case of fecal contamination of groundwater, different water types from the same groundwater area have similar Bacteroidales species. Fourth, the quantitative assays for Bacteroidales 16S rRNA genes commonly used to detect fecal contamination (3, 4, 12, 16, 19, 20, 22, 33, 34) detected Bacteroidales species in deep groundwater and tap water that have no history of fecal contamination. Fifth, Bacteroidales gene copy numbers were significantly higher in tap water than in raw groundwater, demonstrating an increase or growth of Bacteroidales species during the treatment and/or distribution of drinking water. Finally, human- and bovine-specific Bacteroidales strains were not detected in tap water. Consequently, (quantitative) assays for general Bacteroidales 16S rRNA genes are not suitable to discern fecal contamination in groundwater and unchlorinated drinking water in the Netherlands.Nucleotide sequence accession numbers.The 16S rRNA gene sequences obtained in this study were deposited in the GenBank database under accession numbers . GQ169588 to GQ169609相似文献10.
Evidence for Interspecies Gene Transfer in the Evolution of 2,4-Dichlorophenoxyacetic Acid Degraders
Catherine McGowan Roberta Fulthorpe Alice Wright J. M. Tiedje 《Applied and environmental microbiology》1998,64(10):4089-4092
Small-subunit ribosomal DNA (SSU rDNA) from 20 phenotypically distinct strains of 2,4-dichlorophenoxyacetic acid (2,4-D)-degrading bacteria was partially sequenced, yielding 18 unique strains belonging to members of the alpha, beta, and gamma subgroups of the class Proteobacteria. To understand the origin of 2,4-D degradation in this diverse collection, the first gene in the 2,4-D pathway, tfdA, was sequenced. The sequences fell into three unique classes found in various members of the beta and gamma subgroups of Proteobacteria. None of the α-Proteobacteria yielded tfdA PCR products. A comparison of the dendrogram of the tfdA genes with that of the SSU rDNA genes demonstrated incongruency in phylogenies, and hence 2,4-D degradation must have originated from gene transfer between species. Only those strains with tfdA sequences highly similar to the tfdA sequence of strain JMP134 (tfdA class I) transferred all the 2,4-D genes and conferred the 2,4-D degradation phenotype to a Burkholderia cepacia recipient.Bacteria capable of mineralizing 2,4-dichlorophenoxyacetic acid (2,4-D), a commonly used herbicide, are found in many different phylogenetic groups (2, 3, 7, 11, 22, 23). Evidence suggests that numerous variants of 2,4-D catabolic genes exist and that catabolic operons consist of a near-random mixing of these variants (7). Interspecies gene transfer is a well-documented phenomenon (13), and horizontal gene transfer of the 2,4-D-degrading plasmid pJP4 has been shown (3, 5). However, not all 2,4-D catabolic operons are found on plasmids (10, 11, 16, 20). The extent to which other 2,4-D genes have been exchanged in nature is unknown. The aim of this research was to assess the role of horizontal gene transfer in the evolution of 2,4-D-degrading strains. This article summarizes the results of two aspects of this work—the study of the transfer of the entire 2,4-D pathway by using standard mating experiments and a phylogenetic study of the tfdA gene. The tfdA gene codes for an α-ketoglutarate-dependent 2,4-D dioxygenase which converts 2,4-D into 2,4-dichlorophenol and glyoxylate (6). This 861-bp gene was first sequenced from Ralstonia eutropha JMP134 (19). Two more tfdA genes were cloned from chromosomal locations in Burkholderia strain RASC and Burkholderia strain TFD6 (16, 20). These proved to be identical to each other and 78.5% similar to the original. An alignment of the two variants allowed conserved areas to be identified and primers to be designed for the amplification of tfdA-like genes from other sources (24). Sequence analysis of putative tfdA fragments and the small-subunit ribosomal DNA (SSU rDNA) of the strains carrying them allowed us to construct phylogenies of the genes and their hosts and to look for congruency between them.
Open in a separate windowaThe generus and/or species most similar to the strain is given based on similarities of SSU rDNA sequences. bSymbols: +, able to transfer 2,4-D degradation to B. cepacia D5; (+), able to transfer at very low frequency; −, no transfer detected. cND, not determined. d—, no amplificate was obtained. The disappearance of 2,4-D from the culture medium was monitored by high-performance liquid chromatography. Cells were removed by centrifugation, and the supernatant was filtered through 0.2-μm-pore-size filters. These samples were then analyzed on a Lichrosorb Rp-18 column (Anspec Co., Ann Arbor, Mich.) with 60% methanol–40% 0.1% H3PO4 as the eluant. 2,4-D was detected by measuring light absorption at 230 nm. The presence of tfd genes was detected by hybridizing colony blots with a DNA probe derived from the entire pJP4 plasmid. The identity of the colonies was confirmed by probing with the nptII gene of Tn5 (found in B. cepacia D5). Probes were labeled with random hexanucleotides incorporating [32P]dCTP (3,000 Ci/mmol; New England Nuclear, Boston, Mass.). Hybridizations were done under high-stringency conditions by using 50% formamide and Denhardt’s solution (18) at 42°C. Of the 15 unique strains tested, 9 transferred 2,4-D degradation abilities to D5. This transfer was confirmed by hybridization with pJP4 for eight of these strains. B. cepacia RASC could transfer degradative abilities, but neither it nor the transconjugant hybridized to the pJP4 probe. Work subsequent to this study has confirmed that the genes carried by RASC do not hybridize to those found on pJP4 under high-stringency conditions (7).
Mating experiments.
A collection of 2,4-D degraders containing 15 unique strains as determined by genomic fingerprinting (7) was used as a source of donors in a series of mating experiments (Table (Table1).1). Burkholderia cepacia D5, lacking the ability to grow on 2,4-D and not hybridizing to any tfd genes, was used as a recipient in mating experiments. Strain D5 contains neomycin phosphotransferase genes (nptII) carried on transposon Tn5 and is resistant to 50 μg each of kanamycin, carbenicillin, and bacitracin per ml. All of the 2,4-D strains used were sensitive to these antibiotics. Filter matings were performed with a donor-to-recipient ratio of 1:10. Colonies which grew on selective medium (500 ppm of 2,4-D in mineral salts agar [MMO] [23] including 50 μg of kanamycin, carbenicillin, and bacitracin per ml) were subjected to further tests. Their ability to catabolize 2,4-D was tested in liquid medium (same composition as that described above).TABLE 1
2,4-D-degrading strains, geographic origins, and GenBank accession numbersStrain | GenBank accession no. (SSU rDNA) | Origin | Most similar to genus and/or speciesa | Transferb | tfdA typec | GenBank accession no. (tfdA gene) | Reference or source |
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JMP134 | AF049542 | Australia | Ralstonia eutropha | + | I | M16730 | 3 |
EML1549 | AF049546 | Oregon | Burkholderia sp. | + | I | 2 | |
TFD39 | AF049539 | Saskatchewan | Burkholderia sp. | + | I | U43197 | 23 |
K712 | AF049543 | Michigan | Burkholderia sp. | + | I | U43276 | 11 |
TFD9 | AF049537 | Saskatchewan | Alcaligenes xylosoxidans | + | I | U43276 | 23 |
TFD41 | AF049541 | Michigan | Ralstonia eutropha | + | I | 23 | |
TFD38 | AF049540 | Michigan | Ralstonia eutropha | + | NDc | 23 | |
TFD23 | AF049536 | Michigan | Rhodoferax fermentans | + | I | U43276 | 23 |
RASC | AF049544 | Oregon | Burkholderia sp. | (+) | II | U25717 | 2 |
TFD6 | AF049546 | Michigan | Burkholderia sp. | − | II | 23 | |
TFD2 | AF049545 | Michigan | Burkholderia sp. | − | II | 23 | |
TFD31 | AF049536 | Saskatchewan | Rhodoferax fermentans | − | III | 23 | |
B6-9 | AF049538 | Ontario | Rhodoferax fermentans | ND | III | U43196 | 9 |
I-18 | U22836 | Oregon | Halomonas sp. | ND | III | U22499 | 15 |
K1443 | AF049531 | Michigan | Sphingomonas sp. | − | —d | 11 | |
2,4-D1 | AF049535 | Montana | Sphingomonas sp. | − | — | R. Sanford | |
B6-5 | AF049533 | Ontario | Sphingomonas sp. | ND | — | 9 | |
B6-10 | AF049534 | Ontario | Sphingomonas sp. | ND | — | 9 | |
EML146 | AF049532 | Oregon | Sphingomonas sp. | − | — | 2 | |
M1 | AF049530 | French Polynesia | Rhodospeudomonas sp. | ND | R. Fulthorpe |
Phylogenetic analyses.
Total genomic DNA was isolated from 20 unique 2,4-D-degrading strains (including all 15 used for mating experiments) grown on 500 ppm of 2,4-D mineral salts medium amended with 50 ppm of yeast extract. SSU rDNA was amplified by using fD1 and rD1 as primers (25). Putative tfdA fragments were amplified by using primers TVU and TVL as previously described (24). PCR products were purified with a Gene Clean kit (Bio 101, La Jolla, Calif.). Sequencing was done with an Applied Biosystems model 373A automatic sequencer (Perkin-Elmer Cetus) by using fluorescently labeled dye termination at the Michigan State University Sequencing Facility. The sequencing primer used for SSU rDNA fragments was 519R (5′ GTA TTA CCG CGG CTG CTG G-3′). For tfdA fragments, the sequencing primers were the same as the amplification primers. GenBank accession numbers for these sequences are given in Table Table11.The SSU rDNA sequences were compared to sequences in GenBank by using the Basic Local Alignment Search Tool (BLAST) (1), and those strains with the highest maximal segment pair scores were retrieved from GenBank and included in the phylogenetic analysis. Sequences were aligned manually with the software SeqEd (Applied Biosystems) and with MacClade (14). Sites where nucleotides were not resolved for all sequences were deleted from the alignment, as were those nucleotides corresponding to the small loop in this region that is absent in the alpha subgroup of the class Proteobacteria. These deletions left 283 unambiguous sites for the construction of the SSU rDNA phylogenies. Phylogenetic trees were constructed by using the neighbor-joining analysis of pairwise Jukes-Cantor distances (4), and the topology was confirmed by using the maximum parsimony method PAUP (21). Desulfomonile tiedjei of the δ-Proteobacteria was used as an outgroup. Bootstrap analysis based on 100 replicates was used to place confidence estimates on the tree. Only bootstrap values of greater than 50 were used.2,4-D degrader diversity.
The 2,4-D degraders in this study were distributed throughout the alpha, beta, and gamma subgroups of the Proteobacteria (Fig. (Fig.1).1). The lack of representation of gram-positive bacteria is likely a reflection of isolation methods, not of the lack of gram-positive 2,4-D degraders. The majority of these strains were members of the beta subgroup of Proteobacteria, five of which were most closely related to the genus Burkholderia, having at least 92% sequence similarity with each other. Three were closely related to Rhodoferax fermentans (close to the class Comamonadaceae), three were related to Ralstonia eutropha, and one was related to Alcaligenes xylosoxidans. TFD39 falls outside any clear cluster. One member of the γ-Proteobacteria, strain I-18, a haloalkaliphile, was found to be closely related to the salt-loving genus Halomonas (15). The remaining six strains all clustered in the alpha branch of Proteobacteria (Fig. (Fig.1).1). Of this subgroup, five were most closely related to the genus Sphingomonas. One member of the α-Proteobacteria, strain M1, which is the most oligotrophic and slow growing of all the strains used in this study, is 97% similar to Rhodopseudomonas palustris. The character of strain M1 correlates well with its phylogenetic placement near the slow-growing genus Bradyrhizobium. Open in a separate windowFIG. 1Neighbor-joining dendrogram (Jukes-Cantor distances) of SSU rDNA from 2,4-D-degrading bacteria (indicated in boldface type) and reference strains (indicated in italic type). Class I (•), class II (▴), and class III (■) types of tfdA genes are indicated. Bootstrap confidence limits (percentages) are indicated above each branch. Scale bar represents a Jukes-Cantor distance of 0.01.tfdA gene fragments.
tfdA gene fragments were successfully amplified and sequenced from 10 strains of β-Proteobacteria and 1 strain of γ-Protobacteria. None of the strains from the α-Proteobacteria gave any amplificates with these primers. These 313 contiguous nucleotides were aligned with additional tfdA sequences from JMP134 and from strain RASC (Fig. (Fig.2).2). Three distinct classes of tfdA gene sequences with slight variations in each class were found. Class I included fragments from JMP134, TFD39, TFD23, K712, and TFD9 that differed from each other by 2 bp at the most. Class I tfdA genes are probably plasmid encoded. All strains with a class I tfdA gene examined so far contained broad-host-range, self-transmissible plasmids containing 2,4-D genes (2, 3, 11, 17). All of the strains with a class I tfdA gene were able to transfer the 2,4-D phenotype in the mating studies reported above. The class II tfdA sequences included identical fragments amplified from RASC, TFD6, and TFD2 which were 76% similar to those in class I. Class III included identical fragments from strains TFD31, B6-9, and I-18 which were 77% similar to class I genes and 80% similar to class II genes. Both class II and III tfdA genes differed from each other and from class I genes in the same nine sites corresponding to the third base pair of the codons. The tfdA phylogenetic tree is a simple one, with three distinct branches that are incongruent with the SSU rDNA-derived phylogeny (Fig. (Fig.3).3). Class I tfdA sequences were found in Burkholderia-like strains, in strains related to the Comamonas-Rhodoferax group, and in the Ralstonia-Acaligenes group, all in the β-Proteobacteria. Class II sequences are less widely distributed, found only in Burkholderia-like branches. However, even in this subgroup, this tfdA variant is found in strains that differ by 7% at the SSU rDNA level (RASC and TFD2). However, the class III sequences were most interesting, being found both in the Comamonas-Rhodoferax group and in a strain of the γ-Proteobacteria, I-18, strains that differ by 24% at the SSU rDNA level. Class III genes have since been found in a collection of randomly isolated non-2,4-D degraders, including gram-positive bacilli, as well as in various gram-negative bacteria, even though the gene is not expressed (10). Open in a separate windowFIG. 2Alignment of 313 nucleotides of internal fragments of tfdA genes from representative strains. Nucleotides identical to tfdA from pJP4 are represented by periods.Open in a separate windowFIG. 3Phylogenetic incongruency of tfdA genes and SSU rDNA from diverse 2,4-D-degrading bacteria. Dendrograms for tfdA and SSU rDNA are indicated. Shading indicates the type of tfdA sequence, either class I, II, or III. Note that branch lengths are not drawn to scale.An interesting result was the detection of two different tfdA gene variants in sibling strains. TFD23 and TFD31 are identical at the ribosomal gene level, but one harbors a class I gene and the other harbors a class III gene. Similarly, TFD6 and EML159 are rRNA siblings that carry a class II and class I gene, respectively.None of the α-Proteobacteria yielded a PCR product when amplified with the conserved tfdA primers. This finding complements our observation that none of these bacteria hybridized to the tfdA gene, even under conditions of low stringency, indicating that any tfdA-like genes in the α-Proteobacteria are likely to be more divergent from the ones sequenced here (7, 11). In addition, none of the Sphingomonas strains in the study hybridized with a whole pJP4 probe, and similarly, no Sphingomonas strains scored positive for transfer of 2,4-D-degrading ability to recipient B. cepacia D5. Together these results suggest a reduced gene flow between members of the α- and β- or γ-Proteobacteria or poor gene expression of β- or γ-derived genes by α-Proteobacteria. Although plasmid pJP4 is a broad-host-range plasmid and has been known to transfer to α-Proteobacteria such as Rhizobium and Agrobacterium species and to γ-Proteobacteria such as Pseudomonas putida, Pseudomonas fluorescens, and Pseudomonas aeruginosa, the 2,4-D pathway is not expressed in these strains of the α- or γ-Proteobacteria (3). Phylogenetically limited expression of plasmid-borne 3-chlorobenzoate-degradative genes has also been noted for the pseudomonads (8). Subsequent studies have found divergent but related sequences for the tfdB and tfdC genes in 2,4-D-degrading Sphingomonas strains (7, 12, 24).With the exceptions of the minor differences within the class I pJP4-like tfdA sequences, there were no intermediate tfdA sequences. The most likely explanation of this is that the rate of horizontal transfer of the tfd genes is high relative to the rate at which mutations can accumulate. Examination of sequences of tfdA genes from a greater variety of organisms may turn up more intermediate variation. 相似文献11.
Many plant species can be induced to flower by responding to stress factors. The short-day plants Pharbitis nil and Perilla frutescens var. crispa flower under long days in response to the stress of poor nutrition or low-intensity light. Grafting experiments using two varieties of P. nil revealed that a transmissible flowering stimulus is involved in stress-induced flowering. The P. nil and P. frutescens plants that were induced to flower by stress reached anthesis, fruited and produced seeds. These seeds germinated, and the progeny of the stressed plants developed normally. Phenylalanine ammonialyase inhibitors inhibited this stress-induced flowering, and the inhibition was overcome by salicylic acid (SA), suggesting that there is an involvement of SA in stress-induced flowering. PnFT2, a P. nil ortholog of the flowering gene FLOWERING LOCUS T (FT) of Arabidopsis thaliana, was expressed when the P. nil plants were induced to flower under poor-nutrition stress conditions, but expression of PnFT1, another ortholog of FT, was not induced, suggesting that PnFT2 is involved in stress-induced flowering.Key words: flowering, stress, phenylalanine ammonia-lyase, salicylic acid, FLOWERING LOCUS T, Pharbitis nil, Perilla frutescensFlowering in many plant species is regulated by environmental factors, such as night-length in photoperiodic flowering and temperature in vernalization. On the other hand, a short-day (SD) plant such as Pharbitis nil (synonym Ipomoea nil) can be induced to flower under long days (LD) when grown under poor-nutrition, low-temperature or high-intensity light conditions.1–9 The flowering induced by these conditions is accompanied by an increase in phenylalanine ammonia-lyase (PAL) activity.10 Taken together, these facts suggest that the flowering induced by these conditions might be regulated by a common mechanism. Poor nutrition, low temperature and high-intensity light can be regarded as stress factors, and PAL activity increases under these stress conditions.11 Accordingly, we assumed that such LD flowering in P. nil might be induced by stress. Non-photoperiodic flowering has also been sporadically reported in several plant species other than P. nil, and a review of these studies suggested that most of the factors responsible for flowering could be regarded as stress. Some examples of these factors are summarized in 12–14
Open in a separate window 相似文献
Table 1
Some cases of stress-induced floweringStress factor | Species | Flowering response | Reference |
high-intensity light | Pharbitis nil | induction | 5 |
low-intensity light | Lemna paucicostata | induction | 29 |
Perilla frutescens var. crispa | induction | 14 | |
ultraviolet C | Arabidopsis thaliana | induction | 23 |
drought | Douglas-fir | induction | 30 |
tropical pasture Legumes | induction | 31 | |
lemon | induction | 32–35 | |
Ipomoea batatas | promotion | 36 | |
poor nutrition | Pharbitis nil | induction | 3, 4, 13 |
Macroptilium atropurpureum | promotion | 37 | |
Cyclamen persicum | promotion | 38 | |
Ipomoea batatas | promotion | 36 | |
Arabidopsis thaliana | induction | 39 | |
poor nitrogen | Lemna paucicostata | induction | 40 |
poor oxygen | Pharbitis nil | induction | 41 |
low temperature | Pharbitis nil | induction | 9, 12 |
high conc. GA4/7 | Douglas-fir | promotion | 42 |
girdling | Douglas-fir | induction | 43 |
root pruning | Citrus sp. | induction | 44 |
Pharbitis nil | induction | 45 | |
mechanical stimulation | Ananas comosus | induction | 46 |
suppression of root elongation | Pharbitis nil | induction | 7 |
12.
John J. Wiens 《PLoS biology》2021,19(8)
The number of species on Earth is highly uncertain. A recent study has suggested that there are less than 2 million prokaryotic species on Earth; this Formal Comment suggests instead that there are more likely hundreds of millions or billions of species, and that the majority of these are bacteria associated with insects and other animals.The number of species on Earth is a fundamental number in science. Yet, estimates of global biodiversity have been highly uncertain. There are presently approximately 1.9 million described species [1]. Estimates of the actual number (both described and undescribed) have ranged from the low millions into the trillions [2,3]. Furthermore, described species richness [1] is dominated by animals (1.3 million; 68%), not bacteria (approximately 10,000 species; 0.5%). Larsen and colleagues [2] summarized evidence suggesting that the majority of species on Earth may be bacteria associated with insect hosts and that bacterial richness may push global biodiversity into the hundreds of millions of species or even low billions.Louca and colleagues [4] (LEA hereafter) have claimed instead that there are only 40,100 host-associated bacterial species among all animal species and 0.8 to 1.6 million prokaryotic species overall (see their “Author summary”). Strangely, they excluded bacterial species associated with animal hosts from their estimates of total prokaryotic diversity and justified this by claiming that the estimates of Larsen and colleagues [2] were “mathematically flawed.” Here, I examine their claims and present new estimates of global biodiversity.Remarkably, all projections by LEA for host-associated bacterial richness were based on an estimate from one ant genus (Cephalotes), an estimate that is demonstrably incorrect by orders of magnitude (S1 Text). Without examining the underlying data [5], LEA estimated only 40 bacterial species among all 130 ant species in this genus. Yet, simply counting the bacterial species among the 25 sampled ant species in that genus reveals 616 unique bacterial species, of which 539 appear to be unique to the genus and 369 each unique to a single ant species (using the standard 97% cutoff for 16S divergence and data from [5]). Thus, there were >500 bacterial species among 25 ant species, not 40 bacterial species among 130 ant species. This mistake was further exacerbated by inexplicably ignoring data from the other 2 insect genera analyzed by Larsen and colleagues [2], thus maximizing the impact of their incorrect estimate for this genus.Their overall estimate of bacterial richness was also strongly influenced by their questionable assumption that all animal genera can share bacterial species (i.e., reducing their estimate of 3 million host-associated bacterial species to only 40,100). They assumed “a conservative overlap of only 0.1% between any two randomly chosen genera” for the number of bacterial species shared between animal genera. No justification was given for this value of 0.1%, nor were any alternative values explored. Furthermore, they implicitly assumed that any bacterial species can be shared between any pair of animal genera, regardless of their phylogeny, habitat, or geographic range. So, for example, a bacterial species that is a gut endosymbiont of a terrestrial herbivorous insect species endemic to Madagascar could somehow be shared with a deep-sea worm in the northern Pacific Ocean. This is ridiculous: there must be a reason why bacterial species are shared among host species and genera (e.g., shared phylogeny, location, diet). For example, broad-scale studies show that sharing of bacteria among insect hosts is associated with both host phylogeny and diet [6].LEA stated “it is known that substantial overlap exists between the microbiota of different host genera and even of distantly related animal taxa.” However, they provided no numbers to justify this “substantial overlap.” In fact, none of the papers they cited as supporting this assumption actually do (S2 Text). For example, one study [7] found 5 bacterial species shared among 5 insect genera utilizing the same type of host plant (cycads). However, LEA do not mention that this study found 1,789 unique bacterial species among just these 5 insect species (or 177 after filtering). This seems inconsistent with their estimate of only 40,100 bacterial species across all animals. In summary, rather than estimating the overlap of bacterial species among host genera, LEA simply made a number up and combined this with unrealistic, unsupported assumptions about overlap. If LEA had considered Cephalotes (which all their estimates were based on), a survey of this genus and related genera [5] found 1,019 bacterial species, with only 77 of the 616 bacterial species in Cephalotes shared with other sampled genera, and the sharing of bacterial species among hosts strongly related to host phylogeny.Numerous surveys of bacterial diversity in insects strongly suggest that there are far more than 40,100 bacterial species among all animals (8] found roughly twice as many bacterial species as those of approximately 30 insect species [5,9], and the study of 218 insect species [6] found >3.5 times as many as the study of 62 insect species. The simple fact that a study found 9,301 bacterial species among only 218 sampled insect species strongly suggests that there are more than 40,100 bacteria among all animals.Table 1Surveys of bacterial diversity among insect species.LEA incorrectly estimated that a genus of 130 ant species (Cephalotes) hosts only 40 bacterial species and subsequently assumed that all animal genera have the same low number of bacterial species. These broad surveys of bacterial species among insects suggest that many insects (including Cephalotes) host much larger numbers of bacterial species.
Open in a separate windowGiven these problems with the estimate of LEA, what is the actual number of bacterial species on Earth? LEA were correct that Larsen and colleagues [2] only estimated the number of species-specific bacteria per insect host species, and those estimates could be wrong. I therefore recalculated those estimates based on more direct counts of species-specific bacteria from the original studies (S3 Text). In 2]. Specifically, Larsen and colleagues [2] projected 0.209 to 5.8 billion species on Earth, of which 66% to 91% are bacteria, whereas I project 0.183 to 4.2 billion, with 58% to 88% bacteria (2] and are explained below. For each scenario, the projected number of species for each group is shown, along with the percentage of the total number of species belonging to that group (note that plants are <0.5% and are rounded down to 0%). In addition to the 4 scenarios, 4 other assumptions were explored. The first 3 involve different estimated numbers of morphologically cryptic arthropod species per morphology-based insect species (from 6 to 2 to 0; for justification, see [2]). These impact the number of animal species, and all downstream estimates for other groups. The final, fourth set of analyses assumes 6 morphologically cryptic arthropod species and that mites host negligible numbers of nematode species. Scenario 1 assumes that all animal species have a full set of bacterial, protist, and fungal endosymbionts, even if they are parasites, but that microsporidian fungi and apicomplexan protists have little or no host-specific bacterial richness. Scenario 2 assumes that symbionts have limited numbers of symbionts themselves (i.e., nematodes have an average of only one host-specific bacterial species) and that microsporidians and apicomplexans have few or no bacterial species. Scenario 3 assumes that all animal species have a full set of symbiont species and that microsporidians and apicomplexans host (on average) as many bacterial species as animal species do. Scenario 4 is identical to Scenario 1, except that it assumes that mites have reduced species richness relative to other arthropods (0.25 mites∶1 other arthropod species). Note that there is an error in Table 3, Scenario 1 in Larsen and colleagues [2]: There should be 27.2 million animal species, not 20.4. The correct number is used here. Archaean species is considered to be limited overall [2], and so is not treated separately.
Open in a separate windowIn summary, the conclusions of LEA are based on an initial estimate of bacterial richness for one genus that was clearly incorrect, combined with a made-up number (and unrealistic assumptions) to estimate overlap of bacterial species among host genera. Reanalyses here suggest that bacterial richness (and the diversity of life) is more likely in the hundreds of millions or billions. 相似文献
Insect group sampled | Insect species sampled | Unique bacterial species found | References |
---|---|---|---|
Ants (Cephalotes and 3 related genera) | 29 | 1,019 | Sanders and colleagues [5] |
Lycaenid butterflies | 31 | 1,156 | Whitaker and colleagues [9] |
Native Hawaiian insects (beetles, flies, true bugs) | 13 | 1,094 | Poff and colleagues [10] |
Various insect orders | 62 | 2,073 | Colman and colleagues [8] |
21 insect orders | 218 | 9,301 | Yun and colleagues [6] |
Scenario 1 | Scenario 2 | Scenario 3 | Scenario 4 | |||||
---|---|---|---|---|---|---|---|---|
Million species | % of total | Million species | % of total | Million species | % of total | Million species | % of total | |
6 cryptic arthropod species | ||||||||
Animals | 163.2 | 9.4 | 163.2 | 13.7 | 163.2 | 3.9 | 102.0 | 9.4 |
Plants | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 |
Fungi | 165.6 | 9.6 | 165.6 | 13.9 | 165.6 | 3.9 | 104.6 | 9.6 |
Protists | 163.2 | 9.4 | 163.2 | 13.7 | 163.2 | 3.9 | 102.0 | 9.4 |
Bacteria | 1,240.3 | 71.6 | 701.8 | 58.8 | 3,721.0 | 88.3 | 775.2 | 71.5 |
Total | 1,732.7 | 1,194.1 | 4,213.3 | 1,084.1 | ||||
2 cryptic arthropod species | ||||||||
Animals | 54.4 | 9.4 | 54.4 | 13.6 | 54.4 | 3.9 | 34.0 | 9.4 |
Plants | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 |
Fungi | 56.8 | 9.8 | 56.8 | 14.2 | 56.8 | 4.0 | 36.4 | 10.0 |
Protists | 54.4 | 9.4 | 54.4 | 13.6 | 54.4 | 3.9 | 34.0 | 9.4 |
Bacteria | 413.4 | 71.4 | 233.9 | 58.5 | 1,240.3 | 88.2 | 258.4 | 71.1 |
Total | 579.4 | 399.9 | 1,406.3 | 363.1 | ||||
0 cryptic arthropod species | ||||||||
Animals | 27.2 | 9.3 | 27.2 | 13.5 | 27.2 | 3.9 | 17.0 | 9.3 |
Plants | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 |
Fungi | 29.6 | 10.2 | 29.6 | 14.7 | 29.6 | 4.2 | 19.4 | 10.6 |
Protists | 27.2 | 9.3 | 27.2 | 13.5 | 27.2 | 3.9 | 17.0 | 9.3 |
Bacteria | 206.7 | 71.0 | 117.0 | 58.1 | 620.2 | 88.0 | 129.2 | 70.6 |
Total | 291.1 | 201.3 | 704.5 | 182.9 | ||||
Mites host limited nematode richness, 6 cryptic arthropod species | ||||||||
Animals | 122.4 | 9.4 | 122.4 | 11.9 | 122.4 | 3.9 | 91.8 | 9.4 |
Plants | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 | 0.3 | 0 |
Fungi | 124.8 | 9.6 | 124.8 | 12.1 | 124.8 | 3.9 | 94.2 | 9.6 |
Protists | 122.4 | 9.4 | 122.4 | 11.9 | 122.4 | 3.9 | 91.8 | 9.4 |
Bacteria | 930.2 | 71.5 | 661.0 | 64.1 | 2,790.7 | 88.3 | 697.7 | 71.5 |
Total | 1,300.2 | 1,030.9 | 3,160.7 | 975.8 |
13.
14.
Pavan Umate 《Plant signaling & behavior》2011,6(3):335-338
The enzymes called lipoxygenases (LOXs) can dioxygenate unsaturated fatty acids, which leads to lipoperoxidation of biological membranes. This process causes synthesis of signaling molecules and also leads to changes in cellular metabolism. LOXs are known to be involved in apoptotic (programmed cell death) pathway, and biotic and abiotic stress responses in plants. Here, the members of LOX gene family in Arabidopsis and rice are identified. The Arabidopsis and rice genomes encode 6 and 14 LOX proteins, respectively, and interestingly, with more LOX genes in rice. The rice LOXs are validated based on protein alignment studies. This is the first report wherein LOXs are identified in rice which may allow better understanding the initiation, progression and effects of apoptosis, and responses to bitoic and abiotic stresses and signaling cascades in plants.Key words: apoptosis, biotic and abiotic stresses, genomics, jasmonic acid, lipidsLipoxygenases (linoleate:oxygen oxidoreductase, EC 1.13.11.-; LOXs) catalyze the conversion of polyunsaturated fatty acids (lipids) into conjugated hydroperoxides. This process is called hydroperoxidation of lipids. LOXs are monomeric, non-heme and non-sulfur, but iron-containing dioxygenases widely expressed in fungi, animal and plant cells, and are known to be absent in prokaryotes. However, a recent finding suggests the existence of LOX-related genomic sequences in bacteria but not in archaea.1 The inflammatory conditions in mammals like bronchial asthama, psoriasis and arthritis are a result of LOXs reactions.2 Further, several clinical conditions like HIV-1 infection,3 disease of kidneys due to the activation of 5-lipoxygenase,4,5 aging of the brain due to neuronal 5-lipoxygenase6 and atherosclerosis7 are mediated by LOXs. In plants, LOXs are involved in response to biotic and abiotic stresses.8 They are involved in germination9 and also in traumatin and jasmonic acid biochemical pathways.10,11 Studies on LOX in rice are conducted to develop novel strategies against insect pests12 in response to wounding and insect attack,13 and on rice bran extracts as functional foods and dietary supplements for control of inflammation and joint health.14 In Arabidopsis, LOXs are studied in response to natural and stress-induced senescence,15 transition to flowering,16 regulation of lateral root development and defense response.17The arachidonic, linoleic and linolenic acids can act as substrates for different LOX isozymes. A hydroperoxy group is added at carbons 5, 12 or 15, when arachidonic acid is the substrate, and so the LOXs are designated as 5-, 12- or 15-lipoxygenases. Sequences are available in the database for plant lipoxygenases (EC:1.13.11.12), mammalian arachidonate 5-lipoxygenase (EC:1.13.11.34), mammalian arachidonate 12-lipoxygenase (EC:1.13.11.31) and mammalian erythroid cell-specific 15-lipoxygenase (EC:1.13.11.33). The prototype member for LOX family, LOX-1 of Glycine max L. (soybean) is a 15-lipoxygenase. The LOX isoforms of soybean (LOX-1, LOX-2, LOX-3a and LOX-3b) are the most characterized of plant LOXs.18 In addition, five vegetative LOXs (VLX-A, -B, -C, -D, -E) are detected in soybean leaves.19 The 3-dimensional structure of soybean LOX-1 has been determined.20,21 LOX-1 was shown to be made of two domains, the N-terminal domain-I which forms a β-barrel of 146 residues, and a C-terminal domain-II of bundle of helices of 693 residues21 (Fig. 1). The iron atom was shown to be at the centre of domain-II bound by four coordinating ligands, of which three are histidine residues.22Open in a separate windowFigure 1Three-dimensional structure of soybean lipoxygenase L-1. The domain I (N-terminal) and domain II (C-terminal) are indicated. The catalytic iron atom is embedded in domain II (PDB ID-1YGE).21This article describes identification of LOX genes in Arabidopsis and rice. The Arabidopsis genome encodes for six LOX proteins23 (www.arabidopsis.org) (Locus Annotation Nomenclature A* B* C* AT1G55020 lipoxygenase 1 (LOX1) LOX1 859 98044.4 5.2049 AT1G17420 lipoxygenase 3 (LOX3) LOX3 919 103725.1 8.0117 AT1G67560 lipoxygenase family protein LOX4 917 104514.6 8.0035 AT1G72520 lipoxygenase, putative LOX6 926 104813.1 7.5213 AT3G22400 lipoxygenase 5 (LOX5) LOX5 886 101058.8 6.6033 AT3G45140 lipoxygenase 2 (LOX2) LOX2 896 102044.7 5.3177