The authors would like to acknowledge Latoya Wiggins and Katelyn Griffin for their assistance in sample acquisition, as well as Laura Lee Rutherford for their assistance in sampling and molecular analyses. We would also like to thank Sarah Owens from Argonne National Lab for microbiomic sample preparation and sequencing. These investigations were supported equally by the Agricultural Research Service, USDA CRIS Projects &#8220;Pathogen Reduction and Processing Parameters in Poultry Processing Systems&#8221; #6612-41420-017-00 and &#8220;Molecular Approaches for the Characterization of Foodborne Pathogens in Poultry&#8221; #6612-32000-059-00.

1. Mechanical Homogenization of Environmental Poultry Production Samples
<ol style="margin-left: 40px;">
	<li>Prior to extraction, set a water bath to 95 &#176;C and allow the water bath time to reach that temperature.</li>
	<li>Weigh out 0.33 g of soil or fecal material into a 2 ml Lysing Matrix E tube.
	<ol>
		<li>Do not exceed 0.33 g of sample in the tube, since this will cause the following solutions to exceed the capacity of the tube.</li>
		<li>Thaw frozen samples to RT prior to weighing.</li>
		<li>In order to analyze a total of 1 g of soil/feces, weigh out 3 replicate 0.33 g samples for each individual environmental sample.</li>
		<li>Store samples at -20 &#176;C within the conical matrix tubes prior to extraction, if needed.</li>
	</ol>
	</li>
	<li>Add 825 &#181;l Sodium Phosphate Buffer and 275 &#181;l of PLS solution to a sample tube. Mix using a vortex for ~15 sec and then centrifuge the samples at 14,000 x g for 5 min.</li>
	<li>Decant the supernatant and add 700 &#181;l of Buffer ASL. Mix using a vortex for 5 sec.
	<ol>
		<li>Ensure that there is headspace (~10% total volume) available in the conical tube at this point. If there is no headspace, the tubes will have a tendency to leak during the next homogenization step which could lead to cross-contamination and/or sample loss.</li>
	</ol>
	</li>
	<li>Place the samples into a FastPrep 24 Instrument, and homogenize the samples at a speed of 6.0 m/sec for 40 sec.</li>
	<li>Centrifuge the homogenized sample at 14,000 x g for 5 min. Transfer the supernatant to a sterile 2 ml microcentrifuge tube.</li>
	<li>To maximize the DNA recovery from the sample, repeat steps 1.4 to 1.6, combining the supernatants into the same sterile, 2 ml microcentrifuge tube.</li>
</ol>
2. Enzymatic Inhibition of Inhibitors from Sample Homogenates
NOTE: This protocol uses the QIAamp DNA Stool Mini Kit.
<ol style="margin-left: 40px;">
	<li>Incubate the supernatant in a 95 &#176;C water bath for 5 min to maximize DNA recovery from any remaining cells within the supernatant.
	<ol>
		<li>Incubate at 70 &#176;C for samples containing mostly Gram-negative organisms. However if Gram-positive organisms are present (which is the case with poultry fecal samples), incubate at 95 &#176;C.</li>
		<li>Use plastic locking clips on the microcentrifuge tubes to ensure that the tubes will not &#8220;pop&#8221; open and potentially lose sample volume as pressure can build up in these sealed microcentrifuge tubes.</li>
	</ol>
	</li>
	<li>Open each microcentrifuge tube to release the pressure, re-cap the microcentrifuge tubes and mix using a vortex for 15 sec.</li>
	<li>Centrifuge the sample at 14,000 x g for 1 min, remove 1.2 ml of the supernatant and place it into a new sterile 2 ml microcentrifuge tube.</li>
	<li>Add 1 InhibitEx tab to each sample, and mix using a vortex until the sample becomes a uniformly white/off-white liquid.
	<ol>
		<li>Avoid touching the InhibitEx tab while placing it into the microcentrifuge tube containing the sample. To accomplish this, place the blister pack containing the tab directly over the open microcentrifuge tube and gently push the tab out of the blister pack and into the microcentrifuge tube.</li>
	</ol>
	</li>
	<li>Incubate the sample for 1 min at RT (~ 25 &#176;C) and centrifuge at 14,000 x g for 5 min.</li>
	<li>Transfer all liquid to a sterile 1.5 ml microcentrifuge tube and centrifuge at 14,000 x g for 5 min.
	<ol>
		<li>Avoid any remaining particulates that may have pelleted at the bottom of the microcentrifuge tube at the end of step 2.5 when transferring the liquid.</li>
	</ol>
	</li>
</ol>
3. Automated DNA Purification Using the QIAcube Robotic Workstation
NOTE: The number of plastic consumables, the arrangement of the sample rotor adapters within the centrifuge, and the required volumes of the buffers/solutions are dependent on the number of samples that are being run.
<ol style="margin-left: 40px;">
	<li>Add elution tubes and filter tubes to the appropriate slots within the rotor adapters. For each sample, add 400 &#181;l to the middle slot of the rotor adapter. Place the rotor adapters in the workstation centrifuge in the correct arrangement according to the number of samples being purified.
	<ol>
		<li>Make sure that all of the microcentrifuge tube lids are properly secured within the rotor adapter since a failure to do so could result in shearing during one of the centrifugation steps of the purification protocol.</li>
	</ol>
	</li>
	<li>Add the required number of 1,000 &#181;l and 200 &#181;l filter-tips to the workstation, and fill the supplied buffer bottles with the required volume of buffers. 
	NOTE: The buffers required for this purification protocol (AL, AW1, AW2, and AE) are all contained within the QIAamp DNA Stool Mini Kit. The user needs to supply the 100% ethanol that is needed for the AW buffers and as a solution used in the purification process.</li>
	<li>Add the required volume of the supplied proteinase K solution into a sterile 1.5 ml microcentrifuge tube and place it into slot A on the workstation. Also, add the required number (equal to the number of samples being purified) of 2 ml safe-lock microcentrifuge sample tubes RB to the shaker section of the workstation.
	<ol>
		<li>Ensure that the lids of the sample tubes are securely placed into the appropriate slots on the workstation, since a failure to do so will result in an error when the machine initially scans the workstation to make sure all needed plastics and liquids are available for the requested run.</li>
	</ol>
	</li>
	<li>Using the touchscreen on the workstation, select the DNA Stool &#8211; Human Stool &#8211; Pathogen Detection Protocol, and read through the subsequent screens to ensure that the workstation was loaded correctly. Once all check screens are passed, select Start to run this protocol.
	<ol>
		<li>If extracting DNA from more than 12 samples, begin the homogenization process (Step 1) for the next set of samples, since a run of 12 samples takes ~72 min to complete on the workstation.</li>
	</ol>
	</li>
	<li>Remove the samples from the rotor adapters, cap them, and place at -20 &#176;C until needed for subsequent downstream analyses.
	<ol>
		<li>At this point, combine the 3 replicate purifications for an individual sample (total analyzed amount = 1 g) using a centrifugation/evaporation-based system. Combine the replicates and re-elute to a final volume of 100 &#181;l of Tris-ETDA buffer.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The DNA extraction method used effected the quantitative and qualitative total bacterial community estimates for both the fecal and litter samples, supporting the sample analyses dependent nature of DNA extraction methods seen previously1,3,6. For both the fecal and litter samples, the order of performance of the DNA extraction methods was different for the quantitative (Mechanical &#62; Hybrid &#62; Enzymatic) and the qualitative (Enzymatic &#62; Hybrid &#62; Mechanical) total bacterial community estimates. While the hybrid method did not produce the highest quantitative or qualitative estimates, in both cases the hybrid method produced results that were statistically similar to the highest performing extraction method (Figure 1, Table 2), thereby producing the best combination of qualitative and quantitative total bacterial community estimates of the three extraction methods tested. It should ne noted that bacterial community profiles can be strongly effected by DNA extraction method, as seen in the results from this study (Figure 2, Table 1) and other previous studies using environmental samples4,5,8,9,11,12, and these differential effects may strongly influence the analysis and interpretation of the study data. Knowing this, researchers must strongly consider the appropriate extraction method for their sample types and experimental goals on a study-by-study basis.
A key step to the effectiveness of the novel hybrid method is the powerful homogenization step to greatly aid in not only the release of bacterial cells from the complex fecal/soil matrices, but also to break down the tougher cell walls of Gram-positive community members. The higher quantitative estimates (Figure 1) and the higher proportion of Gram-positive Actinobacteria and Firmicutes within the microbiomes for both extraction methods that used this homogenization step (Table 1) are both excellent examples of the effectiveness of this homogenization step compared to the enzymatic method. These results are in agreement with previous studies using fecal samples that have shown the improvement of DNA quantity and improved community analyses when a mechanical homogenization step was included1,2,4-6,9. Once the cells and/or DNA are disassociated/released from these matrices, the use of the &#8220;gold standard&#8221; enzymatic methodology in the hybrid method efficiently purifies the DNA from these samples. The inclusion of the inhibitor-binding step from the enzymatic method within the hybrid extraction protocol allows for a more efficient removal of the inhibitors present within the complex environmental samples used in this study. This step that is absent from the mechanical extraction protocol. Therefore, the hybrid extraction method, with the addition of a mechanical homogenization step in conjunction with the enzymatic purification of the released DNA, is an effective method for the assessment of total bacterial communities from environmental poultry production samples.
This method, as described, contains some specific requirements of which users will need to be aware. First is the use of a high-powered homogenizer as well as some components of a commercially available extraction kit. Previous studies demonstrated that other mechanical bead-beating steps or high-powered homogenizers improve DNA extraction efficiency and yield4,5,9; therefore, a different type of mechanical homogenization step could be integrated into the hybrid extraction method (although the use of different specialized equipment may be required). The hybrid method described here also uses the bead-beating tubes that are specifically designed for the homogenizer and specific for environmental samples due to the success of this lysing matrix in previous community-based studies from poultry samples15,31-33. Finally, the &#8220;gold standard&#8221; enzymatic DNA extraction method (QIAamp DNA Stool Mini Kit) is required for the enzymatic purification of the sample DNA in the hybrid protocol, since it has been shown to be the most effective enzymatic extraction method when using fecal samples3,18,19. While the use of the robotic workstation is not required (instructions with the kit can be performed by hand), the use of the robotic workstation greatly increases throughput, minimizes processing error, and results in higher quality DNA for further downstream analysis7. Automation of this type, using the existing mechanical kits is not currently available, which represents another advantage of the hybrid method.
Currently, the robotic workstation described in this protocol is limited to processing a maximum of 12 samples in 72 min, but preliminary work has shown that this method works equally well in the higher throughput workstation that can process 96 samples within 40 min&#160;(data not shown). Further testing of this much higher throughput option will greatly enhance the efficacy of the hybrid method since more samples could be processed in a semi-automated manner within a typical day. Also, preliminary data using this novel DNA extraction method has shown that it is effective for other poultry-related samples (e.g., skin/feather rinses, carcass rinses, cecal content homogenates) as well as other types of environmental samples (e.g., agricultural soils, swine feces, horse feces, goat feces, bovine feces). Therefore, future studies will need to determine the efficacy of this hybrid method for the extraction of DNA from many agricultural and environmental samples. Further validation of this method by future studies would potentially allow for a wider adoption of this method for the assessment of total bacterial communities from a variety of agricultural and environmental settings.

When analyzing complex clinical or environmental samples (e.g., feces, soils), there are two main methodologies used for the extraction of DNA. The first is a mechanical disruption of the matrix using an intense bead-beating step, while the second is an enzymatic disruption of the matrix to chemically release bacterial cells and inhibit PCR inhibitors from the matrix simultaneously. Given the different means by which these two types of extraction methods work, it is not surprising that previous studies demonstrated that the appropriate DNA extraction method is both highly sample and analysis dependent. Comparative DNA extraction studies previously showed that some methods are more appropriate for improved DNA quality and quantity from environmental samples1-3, while others demonstrated improvements for community-level analyses such as denaturing gradient gel electrophoresis (DGGE)4-6, terminal restriction fragment length polymorphism (T-RFLP)7, automated ribosomal intergenic spacer analysis (ARISA)8, and phylogenetic microarrays9. Therefore, appropriate DNA extraction methods need to be used, or developed, according to the types of environmental samples and the types of analyses being performed on those samples, especially given the recent advancements in bacterial community analyses. 
Next generation sequencing, in conjunction with more quantitative community assessments (e.g., quantitative PCR (qPCR)), is becoming more prevalent in the environmental and clinical sciences, however, very little research has been performed to determine the effect of DNA extraction methods on these data sets. Most DNA extraction comparison studies dealt with microbiomic community estimates from human or human model samples10,11, not agricultural animal samples. The few poultry-focused next generation sequencing studies dealt with specific metagenomic12,13 or microbiomic14 questions; they did not discuss the effect of DNA extraction method on the resulting microbiomic analyses. Considering the complex nature of environmental samples related to poultry production (e.g., feces, litter/bedding, pasture soil), DNA extraction methods need to be carefully selected. Poultry-related environmental samples are known to contain large numbers of PCR inhibitors and up to 500-fold DNA extract dilutions have been required for PCR and subsequent downstream analysis15-17. Therefore it is essential that DNA extraction methods be optimized for these types of samples in order to not only physically disrupt the matrix, but also to be able to reduce/eliminate the large number of inhibitors that are present. 
The QIAamp DNA Stool Mini Kit, an enzymatic extraction method, has been considered the &#8220;gold standard&#8221; when extracting DNA from difficult gut/fecal samples1,18,19 and has been applied successfully to poultry environmental samples8,14. The enzymatic removal of PCR inhibitors through the use of a proprietary matrix is one of the greatest advantages of using this method for these types of environmental samples, as is the ability to significantly improve throughput (and reduce sample processing error) using automated workstations. One major disadvantage is the lack of a mechanical homogenization step to physically disassociate bacterial cells from the environmental matrix. When testing gut and fecal samples of non-poultry origin, the addition of a bead-beating or mechanical disruption step within a DNA extraction protocol significantly increased extraction efficiency9, DNA yield/quality1,4,5 and significantly improved downstream community analyses in terms of richness, diversity, and coverage5,6,11. These studies compared not only mechanical bead-beating methods to the &#8220;gold standard&#8221; enzymatic method, but some also added the mechanical bead-beating step to the enzymatic protocol to improve results6,9,11. 
According to the results from the above studies, bacterial community analyses (both qualitative and quantitative) could be improved from poultry-related environmental samples through the addition of a mechanical homogenization step to the enzymatic method. Therefore, the goal of this study was twofold: (1) to develop a novel DNA extraction technique that utilizes the most desirable aspects of both the mechanical (powerful homogenization step) and enzymatic (PCR inhibitor removal and automation) extraction methods and (2) compare the quantitative (via qPCR) and qualitative (via microbiomics) bacterial community assessments of this novel method to representative mechanical and enzymatic methods.

<table><tbody><tr><td>Lysing Matrix E tube</td><td>MPBio</td><td>6914-050</td><td>Different sizes available and the last 3 numbers of the cat. No. indicate size (-050 = 50 tubes, -200 = 200 tubes, -1000 = 1,000 tubes)</td></tr><tr><td>Sodium Phosphate Solution</td><td>MPBio</td><td>6570-205</td><td>Can be purchased individually, or also contained within the FastDNA Spin Kit for feces (Cat. No. 116570200)</td></tr><tr><td>PLS Buffer</td><td>MPBio</td><td>6570-201</td><td></td></tr><tr><td>Buffer ASL (560 ml)</td><td>Qiagen</td><td>19082</td><td></td></tr><tr><td>FastPrep 24 homogenizer</td><td>MPBio</td><td>116004500</td><td>48 x 2 ml HiPrep adapter (Cat. No. 116002527) available to double throughput of mechanical homogenization step</td></tr><tr><td>QIAamp DNA Stool Mini Kit</td><td>Qiagen</td><td>51504</td><td></td></tr><tr><td>QIAcube24 (110V)</td><td>Qiagen</td><td>9001292</td><td>Preliminary results show that QIAcube HT (Cat. No. 9001793) can be used to improve throughput, but different consumables are required of this machine and more comparative work needs to be done.</td></tr><tr><td>Filter-Tips, 1,000 ml (1024)</td><td>Qiagen</td><td>990352</td><td></td></tr><tr><td>Filter-Tips, 200 ml (1024)</td><td>Qiagen</td><td>990332</td><td></td></tr><tr><td>QIAcube Rotor Adapters (10 x 24)</td><td>Qiagen</td><td>990394</td><td>For 1.5 ml microcentrifuge tubes included with in the rotor adapter kit there is an alternative.&amp;nbsp; It is Sarstedt Micro tube 1.5 ml Safety Cap, Cat. No. 72.690</td></tr><tr><td>Sample Tubes RB (2 ml)</td><td>Qiagen</td><td>990381</td><td>Alternative: Eppendorf Safe-Lok micro test tube, Cat. No. 022363352</td></tr></tbody></table>

a hybrid dna extraction method for the qualitative and quantitative assessment of bacterial communities from poultry production samples

The efficacy of DNA extraction protocols can be highly dependent upon both the type of sample being investigated and the types of downstream analyses performed. Considering that the use of new bacterial community analysis techniques (e.g., microbiomics, metagenomics) is becoming more prevalent in the agricultural and environmental sciences and many environmental samples within these disciplines can be physiochemically and microbiologically unique (e.g., fecal and litter/bedding samples from the poultry production spectrum), appropriate and effective DNA extraction methods need to be carefully chosen. Therefore, a novel semi-automated hybrid DNA extraction method was developed specifically for use with environmental poultry production samples. This method is a combination of the two major types of DNA extraction: mechanical and enzymatic. A two-step intense mechanical homogenization step (using bead-beating specifically formulated for environmental samples) was added to the beginning of the &#8220;gold standard&#8221; enzymatic DNA extraction method for fecal samples to enhance the removal of bacteria and DNA from the sample matrix and improve the recovery of Gram-positive bacterial community members. Once the enzymatic extraction portion of the hybrid method was initiated, the remaining purification process was automated using a robotic workstation to increase sample throughput and decrease sample processing error. In comparison to the strict mechanical and enzymatic DNA extraction methods, this novel hybrid method provided the best overall combined performance when considering quantitative (using 16S rRNA qPCR) and qualitative (using microbiomics) estimates of the total bacterial communities when processing poultry feces and litter samples.

For this study, fresh fecal droppings and litter samples were recovered from a commercial broiler house (~25,000 birds) in the southeastern US. The broilers (Gallus gallus) were Cobb-500 crosses, and they were 59 days old at the time of sampling. Fresh fecal and litter samples were recovered from four distinct areas within the house (near cooling pad, near the waterer/feeder lines, in between the waterer/feeder lines, and near exhaust fans), and samples from each of these areas contained five pooled samples from within that area. Samples from the four areas of the house were extracted and quantitatively/qualitatively analyzed separately according to sample type (fecal or litter). The data presented below represent the mean values for the entire house for both environmental sample types.
Total 16S rDNA abundances, an estimate of the total bacterial community contained within a sample, were determined using a previously published qPCR protocol20, and values were normalized using the amount of DNA recovered from each extraction method (based on fluorometric analysis described previously21. Of the three extraction methods, Figure 1 shows that the mechanical method consistently provided the highest total bacterial normalized gene abundance estimates for both the fecal and litter samples (5.85 &#177; 0.16 and 5.56 &#177; 0.08 log10 16S rDNA copies ng-1 DNA extracted, respectively); whereas, the enzymatic method consistently provided the lowest fecal and litter estimates (4.83 &#177; 0.54 and 4.61 &#177; 0.13 log10 16S rDNA copies ng-1 DNA extracted, respectively). The novel hybrid extraction method yielded normalized 16S rDNA abundances that were found to be significantly (p &#8804; 0.05) higher than the enzymatic method, but statistically similar to the mechanical method for both the fecal and litter samples (5.50 &#177; 0.16 and 5.23 &#177; 0.14 log10 16S rDNA copies ng-1 DNA extracted, respectively). Therefore, both the mechanical and hybrid extraction methods provided a greater qualitative estimate of the total bacterial communities within these samples than the enzymatic method.
In order to qualitatively assess the total bacterial communities from each sample type and extraction method, the microbiomic workflow as reviewed in Navas-Molina et al. (2013)22 was used. In short, the V4 region of the 16S rRNA gene was PCR amplified with primers containing MiSeq sequencing adapters and Golay barcodes and sequenced on the Illumina MiSeq platform23,24. Raw sequencing data was then processed in QIIME 1.7.0-dev25,26,27 using the default parameters. Sequence data processing included: quality-filtering; OTU clustering (open-reference at 97% threshold) using UCLUST27; OTU abundance filtering level of &#60;0.005%22,31; taxonomy assignment using the RDP Classifier against the Greengenes 13_5 reference database28; sequence alignment with PyNAST29 against the Greengenes core set28; phylogenetic tree construction using FastTree30. The core_diversity_analyses.py script was used to run all alpha beta diversity metrics and to generate all plots, charts, and statistics at a sequencing depth of 7865 sequences per sample.
The chosen DNA extraction method exhibited a clear influence on the recovered fecal and litter microbiomes. Starting at the phylum-level (Table 1), the overall data (accounting for both fecal and litter samples) revealed that methods possessing the highly disruptive homogenization step (mechanical, hybrid) typically yielded communities with higher abundances of Gram-positive phyla (98.0 and 97.49%, respectively) relative to the enzymatic method (81.74%). Conversely, Gram-negative phyla represented a much greater part of the overall bacterial community recovered from the enzymatic method (17.49%) compared to either the mechanical (1.89%) or hybrid (2.38%) methods. This differential extraction effect on the overall microbiomic profiles and the prevalence of Gram-positive and Gram-negative organisms was effectively demonstrated at the genus level (Figure 2).The mechanical and hybrid methods produced relatively similar microbiomic profiles, while the enzymatic method microbiome showed a different distribution of the relative abundance of the recovered taxa. For example, Gram-negative Alistipes spp.in the feces samples (the red block with the A; 7.71% of the total community) extracted using the enzymatic method is severely reduced in the fingerprints for the other two extraction methods (red arrows; 0.47-0.81%), but the Gram-positive Lactobacillus spp. (large purple block with the L) was much more prevalent is reduced when using the enzymatic method (42.71%) as compared to the mechanical or hybrid methods (57.22% and 61.85%, respectively). Nonetheless, while the relative abundances of the taxa exhibited some differences between the three extraction methods, the total number of taxa discovered were similar for all three methods for both the fecal (92 of 112 taxa) and litter (96 of 112 taxa).
The enzymatic method consistently yielded bacterial communities with the greatest richness, diversity, and evenness (using the chao1, Phylogenetic Diversity, and equitability metrics, respectively) for both the fecal and litter samples, with the mechanical method consistently showing the lowest estimates for these ecological parameters (Table 2). The hybrid method consistently demonstrated ecological metrics intermediary to the other two methods. Statistically, the ecological estimates from all samples, regardless of extraction method, were found to be similar, although enzymatic method did result in a significantly richer (p = 0.045) microbiome compared to the mechanical method when analyzing the litter samples. While no other significant differences were found between extraction methods, the greatest differences between the ecological parameters were typically found between the enzymatic and mechanical methods, with the hybrid method performing equally well to the other two methods; similar to the quantitative assessment of these extraction methods (Figure 1).
<img alt="Figure 1" src="/files/ftp_upload/52161/52161fig1highres.jpg" /> 
Figure 1.&#160;Qualitative bacterial community comparison based on DNA extraction methods. Columns represent the mean log10-transformmed 16S rRNA gene copies as determined by qPCR normalized against the amount of DNA recovered from fecal (solid bars) or litter (dashed bars) sample (based on fluorometric determination). The DNA extraction method used to obtain these values is indicated on the x-axis. The error bars represent the standard deviation between 4&#160;replicate samples. For each sample type (Feces or Litter), the letters above the columns represent significantly different means based on one-way ANOVA analysis using the Tukey&#8217;s multiple comparison test (p = 0.05).
<img alt="Figure 2" src="/files/ftp_upload/52161/52161fig2highres.jpg" /> 
Figure 2.&#160;Genus-level microbiomic comparison of the DNA extraction methods. Each column represents the average total bacterial community (based on 16S rRNA gene Illumina MiSeq data) of recovered taxa for each extraction method for the feces (first three columns) and litter (last three columns) samples. Each column represents the average of four different samples from distinct areas within the broiler house. The letter at the base of the columns indicates the extraction method (M = Mechanical, E = Enzymatic, H = Hybrid). The different colors in each column represent the relative abundance of a genus within the overall community, and color for each genera is the same for all columns.
<img alt="Table 1" fo:content-width="6in" src="/files/ftp_upload/52161/52161table1.jpg" width="600" /> 
Table 1. Phylum-level microbiomic comparison of DNA extraction methods. Relative abundances (% total recovered sequences) for each DNA extraction method of the 2 Gram-positive (Actinomycetes and Firmicutes) and 3 Gram-negative (Bacteriodetes, Proteobacteria, Tenericutes) phyla recovered from the microbiomic analysis from both samples types.
<img alt="Table 2" fo:content-width="7in" src="/files/ftp_upload/52161/52161table2.jpg" width="700" /> 
Table 2. Pairwise comparisons of richness, diversity,&#160;and evenness estimates for the three DNA extraction methods based on genus-level microbiomic bacterial community analyses using QIIME. For both sample types, pairwise comparisons were performed for the three possible extraction method combinations. The total bacterial community parameters assessed include richness (&#945;-diversity based on the chao1 statistic), diversity (&#946;-diversity based on the Phylogenetic Diversity statistic), and evenness (based on the equitability statistic). Values represent the mean (standard deviation) of 4&#160;distinct area samples within the broiler house, and an &#945; = 0.05 level was used to determine significance between pairwise comparisons. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/52161/52161table2.jpg" target="_blank">Please click here to view a larger version of this table.</a>

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A Hybrid DNA Extraction Method for the Qualitative and Quantitative Assessment of Bacterial Communities from Poultry Production Samples

A novel semi-automated hybrid DNA extraction method for use with environmental poultry production samples was developed and demonstrated improvements over a common mechanical and enzymatic extraction method in terms of the quantitative and qualitative estimates of the total bacterial communities.

The efficacy of DNA extraction protocols can be highly dependent upon both the type of sample being investigated and the types of downstream analyses ...

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Nanyang Technological University

Research

JoVE Journal

Biology

14.5K Views.  USDA-Agricultural Research Service. A novel semi-automated hybrid DNA extraction method for use with environmental poultry production samples was developed and demonstrated improvements over a common mechanical and enzymatic extraction method in terms of the quantitative and qualitative estimates of the total bacterial communities. 

Video: A Hybrid DNA Extraction Method for the Qualitative and Quantitative Assessment of Bacterial Communities from Poultry Production Samples

Hybrid De Novo Genome Assembly for the Generation of Complete Genomes of Urinary Bacteria using Short- and Long-read Sequencing Technologies

Complete genome sequences provide valuable data for the understanding of genetic diversity and unique colonization factors of urinary microbes. These data may include mobile genetic elements, such as plasmids and extrachromosomal phage, that contribute to the dissemination of antimicrobial resistance and further complicate treatment of urinary tract infection (UTI). In addition to providing fine resolution of genome structure, complete, closed genomes allow for the detailed comparative genomics and evolutionary analyses. The generation of complete genomes de novo has long been a challenging task due to limitations of available sequencing technology. Paired-end Next Generation Sequencing (NGS) produces high quality short reads often resulting in accurate but fragmented genome assemblies. On the contrary, Nanopore sequencing provides long reads of lower quality normally leading to error-prone complete assemblies. Such errors may hamper genome-wide association studies or provide misleading variant analysis results. Therefore, hybrid approaches combining both short and long reads have emerged as reliable methods to achieve highly accurate closed bacterial genomes. Reported herein is a comprehensive method for the culture of diverse urinary bacteria, species identification by 16S rRNA gene sequencing, extraction of genomic DNA (gDNA), and generation of short and long reads by NGS and Nanopore platforms, respectively. Additionally, this method describes a bioinformatic pipeline of quality control, assembly, and gene prediction algorithms for the generation of annotated complete genome sequences. Combination of bioinformatic tools enables the selection of high quality read data for hybrid genome assembly and downstream analysis. The streamlined approach for the hybrid de novo genome assembly described in this protocol may be adapted for the use in any culturable bacteria.

Hybrid De Novo Genome Assembly for the Generation of Complete Genomes of Urinary Bacteria using Short- and Long-read Sequencing Technologies

This protocol details a comprehensive approach for the culturing, sequencing, and de novo hybrid genome assembly of urinary bacteria. It provides a reproducible procedure for the generation of complete, circular genome sequences useful in studying both chromosomal and extrachromosomal genetic elements contributing to urinary colonization, pathogenesis, and antimicrobial resistance dissemination.

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Genetics

Isolation, Propagation, and Identification of Bacterial Species with Hydrocarbon Metabolizing Properties from Aquatic Habitats

Hydrocarbon pollutants are recalcitrant to degradation and their accumulation in the environment is toxic to all life forms. Bacteria encode numerous catalytic enzymes and are naturally capable of metabolizing hydrocarbons. Scientists harness biodiversity in aquatic ecosystems to isolate bacteria with biodegradation and bioremediation potential. Such isolates from the environment provide a rich set of metabolic pathways and enzymes, which can be further utilized to scale up the degradation process at an industrial scale. In this article, we outline the general process of isolation, propagation, and identification of bacterial species from aquatic habitats and screen their ability to utilize hydrocarbons as the sole carbon source in vitro using simple techniques. The present protocol describes the isolation of various bacterial species and their subsequent identification using the 16S rRNA analysis. The protocol also presents steps for characterizing the hydrocarbon degrading potential of bacterial isolates. This protocol will be useful for researchers trying to isolate bacterial species from environmental habitats for their biotechnological applications.

We present the process of isolating, propagating, and characterizing hydrocarbon-degrading bacteria from aquatic habitats. The protocol outlines bacterial isolation, identification by the 16S rRNA method, and testing of their hydrocarbon-degrading potential. This article would help researchers in characterizing microbial biodiversity in environmental samples, and specifically screen for microbes with bioremediation potential.

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Extraction of High Molecular Weight DNA from Microbial Mats

Successful and accurate analysis and interpretation of metagenomic data is dependent upon the efficient extraction of high-quality, high molecular weight (HMW) community DNA. However, environmental mat samples often pose difficulties to obtaining large concentrations of high-quality, HMW DNA. Hypersaline microbial mats contain high amounts of extracellular polymeric substances (EPS)1 and salts that may inhibit downstream applications of extracted DNA. Direct and harsh methods are often used in DNA extraction from refractory samples. These methods are typically used because the EPS in mats, an adhesive matrix, binds DNA2,3 during direct lysis. As a result of harsher extraction methods, DNA becomes fragmented into small sizes4,5,6.
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We provide an improved protocol for extracting high molecular weight DNA from hypersaline microbial mats. Microbial cells are separated from the mat matrix prior to DNA extraction and purification. This enhances the concentrations, quality, and size of the DNA. The protocol may be used for other refractory samples.

Successful and accurate analysis and interpretation of metagenomic data is dependent upon the efficient extraction  of high-quality, high molecular weight ...

Production of Germ-Free Fast-Growing Broilers from a Commercial Line for Microbiota Studies

Studies of the gut microbiota contribution to the host physiology and immunocompetence are facilitated by the availability of germ-free animal models, which are considered the gold standard. Nesting birds are ideal models for the production of germ-free animals since there is no need to raise their relatives under sterile conditions. Germ-free chickens are mainly generated from specific-pathogen-free (SPF) experimental lines, which are poorly representative of commercial chicken lines.&#160;The method proposed here allowed the production of germ-free chickens from the fast growing broiler line Ross PM3, commonly used by the poultry industry.&#160;Eggs were quickly collected after laying at a broiler breeder farm.&#160;They underwent a strict decontamination process from the collection to the introduction in a sterile egg hatching isolator.&#160;The chicks have been hatched and kept in these sterile isolators during the period necessary to control their sterility. Originally developed for an experimental SPF white leghorn line, the present protocol has been adapted not only to the Ross PM3 broiler line but also to quails. It therefore represents a robust and readily adaptable procedure to other poultry species and nesting birds of economic, biological or ecological relevance.

Here we describe a method for the generation of germ-free chicks from eggs of a commercial broiler line, the Ross PM3. This method can be adapted for the generation of germ-free animals from other poultry species.

Studies of the gut microbiota contribution to the host physiology and immunocompetence are facilitated by the availability of germ-free animal models, ...

Immunology and Infection

Who is Who? Non-invasive Methods to Individually Sex and Mark Altricial Chicks

Many experiments require early determination of offspring's sex as well as early marking of newborns for individual recognition. According to animal welfare guidelines, non-invasive techniques should be preferred whenever applicable. In our group, we work on different species of song birds in the lab and in the field, and we successfully apply non-invasive methods to sex and individually mark chicks. This paper presents a comprehensive non-invasive tool-box. Sexing birds prior to the expression of secondary sexual traits requires the collection of DNA-bearing material for PCR. We established a quick and easy method to sex birds of any age (post hatching) by extracting DNA from buccal swabs. Results can be obtained within 3 hours. For individual marking chick's down feathers are trimmed in specific patterns allowing fast identification within the hatching order. This set of methods is easily applicable in a standard equipped lab and especially suitable for working in the field as no special equipment is required for sampling and storage. Handling of chicks is minimized and marking and sexing techniques are non-invasive thereby supporting the RRR-principle of animal welfare guidelines.

This protocol provides a convenient set of methods, which enables extremely fast, easy, non-invasive, reliable and low-cost, molecular sex determination of birds and their non-invasive, quick, safe and easily recognizable marking shortly after hatching. Only limited handling of chicks is required. This convenient toolbox of methods complies entirely with the RRR-guidelines.

Many experiments require early determination of offspring's sex as well as early marking of newborns for individual recognition. According to animal ...

Efficient Nucleic Acid Extraction and 16S rRNA Gene Sequencing for Bacterial Community Characterization

There is a growing appreciation for the role of microbial communities as critical modulators of human health and disease. High throughput sequencing technologies have allowed for the rapid and efficient characterization of bacterial communities using 16S rRNA gene sequencing from a variety of sources. Although readily available tools for 16S rRNA sequence analysis have standardized computational workflows, sample processing for DNA extraction remains a continued source of variability across studies. Here we describe an efficient, robust, and cost effective method for extracting nucleic acid from swabs. We also delineate downstream methods for 16S rRNA gene sequencing, including generation of sequencing libraries, data quality control, and sequence analysis. The workflow can accommodate multiple samples types, including stool and swabs collected from a variety of anatomical locations and host species. Additionally, recovered DNA and RNA can be separated and used for other applications, including whole genome sequencing or RNA-seq. The method described allows for a common processing approach for multiple sample types and accommodates downstream analysis of genomic, metagenomic and transcriptional information.

We describe an efficient, robust, and cost effective method for extracting nucleic acid from swabs for characterization of bacterial communities using 16S rRNA gene amplicon sequencing. The method allows for a common processing approach for multiple sample types and accommodates a number of downstream analytic processes.

There is a growing appreciation for the role of microbial communities as critical modulators of human health and disease. High throughput sequencing ...

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The cryosphere offers access to preserved organisms that persisted under past environmental conditions. In fact, these frozen materials could reflect conditions over vast time periods and investigation of biological materials harbored inside could provide insight of ancient environments. To appropriately analyze these ecosystems and extract meaningful biological information from frozen soils and ice, proper collection and processing of the frozen samples is necessary. This is especially critical for microbial and DNA analyses since the communities present may be so uniquely different from modern ones. Here, a protocol is presented to successfully collect and decontaminate frozen cores. Both the absence of the colonies used to dope the outer surface and exogenous DNA suggest that we successfully decontaminated the frozen cores and that the microorganisms detected were from the material, rather than contamination from drilling or processing the cores.

The cryosphere offers access to preserved organisms that persisted under past environmental conditions. A protocol is presented to collect and decontaminate permafrost cores of soils and ice. The absence of exogenous colonies and DNA suggest that microorganisms detected represent the material, rather than contamination from drilling or processing.

The cryosphere offers access to preserved organisms that persisted under past environmental conditions. In fact, these frozen materials could reflect ...

Fluorescence In Situ Hybridization on DNA Halo Preparations to Reveal Whole Chromosomes, Telomeres and Gene Loci

The genome is associated with several structures inside cell nuclei, in order to regulate its activity and anchor it in specific locations. These structures are collectively known as the nucleoskeleton and include the nuclear lamina, the nucleoli, and nuclear bodies. Although many variants of fluorescence in situ hybridization (FISH) exist to study the genome and its organization, these are often limited by resolution and provide insufficient information on the genome&#39;s association with nuclear structures. The DNA halo method uses high salt concentrations and nonionic detergents to generate DNA loops that remain anchored to structures within nuclei through attachment regions within the genome. Here, soluble nuclear proteins, such as histones, lipids, and DNA not tightly bound to the nuclear matrix, are extracted. This leads to the formation of a halo of unattached DNA surrounding a residual nucleus which itself contains DNA closely associated with internal nuclear structures and extraction-resistant proteins. These extended DNA strands enable increased resolution and can facilitate physical mapping. In combination with FISH, this method has the added advantage of studying genomic interactions with all the structures that the genome is anchored by. This technique, termed HALO-FISH, is highly versatile whereby DNA halos can be coupled with nucleic acid probes to reveal gene loci, whole chromosomes, alpha satellite, telomeres and even RNA. This technique provides an insight into nuclear organization and function in normal&#160;cells and in disease progression such as with cancer.

Combining DNA halo preparations with fluorescence in situ hybridization enables high-resolution analysis of genomic interactions with the nucleoskeleton. Attached genome leads to hybridized fluorescent signals located within the residual extracted nuclei, whereas non-attached genome is in the halo of DNA surrounding the residual nuclei.

The genome is associated with several structures inside cell nuclei, in order to regulate its activity and anchor it in specific locations. These ...

Cultivation Strategies for Uncultured Microbial Diversity in the Chicken GI Tract

Strategies to Enhance Cultivation of Anaerobic Bacteria from Gastrointestinal Tract of Chicken

The gastrointestinal tract (GIT) of chicken is a complex ecosystem harboring trillions of microbes that play a pivotal role in the host&#39;s physiology, digestion, nutrient absorption, immune system maturation, and prevention of pathogen intrusion. For optimal animal health and productivity, it is imperative to characterize these microorganisms and comprehend their role. While the GIT of poultry holds a reservoir of microorganisms with potential probiotic applications, most of the diversity remains unexplored. To enhance our understanding of uncultured microbial diversity, concerted efforts are required to bring these microorganisms into culture. Isolation and cultivation of GIT-colonizing microorganisms yield reproducible material, including cells, DNA, and metabolites, offering new insights into metabolic processes in the environment. Without cultivation, the role of these organisms in their natural settings remains unclear and limited to a descriptive level. Our objective is to implement cultivation strategies aimed at improving the isolation of a diverse range of anaerobic microbes from the chicken&#39;s GIT, leveraging multidisciplinary knowledge from animal physiology, animal nutrition, metagenomics, feed biochemistry, and modern cultivation strategies. Additionally, we aim to implement the use of proper practices for sampling, transportation, and media preparation, which are known to influence isolation success. Appropriate methodologies should ensure a consistent oxygen-free environment, optimal atmospheric conditions, appropriate host incubation temperature, and provision for specific nutritional requirements in alignment with their distinctive needs. By following these methodologies, cultivation will not only yield reproducible results for isolation but will also facilitate isolation procedures, thus fostering a comprehensive understanding of the intricate microbial ecosystem within the chicken&#39;s GIT.

Author Spotlight: Advancing Intestinal Bacteria Cultivation for Poultry

The protocol presents two methodologies to improve the isolation of anaerobic intestinal bacteria. The first focuses on the isolation of a diverse range of bacteria using different culture media. The second focuses on the cultivation steps of a specific microbial group, possibly assimilating myo-inositol, to fully comprehend its ecological significance.

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Modified Most Probable Number Assay to Quantify Salmonella in Raw and Ready-to-Cook Chicken Products

Salmonella is a leading cause of foodborne illness in the United States, particularly in poultry products. Traditional methods for detecting Salmonella focus on prevalence rather than quantification, which limits their utility in assessing contamination levels and risks. This study introduces a novel most probable number (MPN) assay designed to quantify Salmonella in ready-to-cook poultry products, such as chicken cordon bleu. The method involves washing the poultry sample, concentrating the rinse through centrifugation, and serially diluting it in a 48-well block. The MPN assay is integrated with the loop-mediated isothermal amplification (LAMP) method to provide a sensitive, accurate, and rapid quantification of Salmonella contamination within the same timeframe as existing Food Safety and Inspection Service (FSIS) protocols. Results show a strong linear correlation between the MPN-LAMP measurements and theoretical inoculation levels (R&#178; = 0.933). However, variability at lower concentrations highlights challenges in accurately detecting Salmonella at these levels, with the practical lower limit of detection estimated at approximately 300 CFU/g. Potential refinements to improve the protocol's applicability include increasing the quantity sampled to further improve the limit of detection, optimizing enrichment media formulations, and expanding molecular detection to target multiple Salmonella serovars. Overall, this study presents a practical tool for the food industry, enabling reliable quantification of Salmonella contamination in poultry products, contributing to improved food safety and public health.

Accurately quantifying Salmonella in poultry at low levels is a current industrial and regulatory challenge. This protocol describes an MPN assay that enables quantification of Salmonella in raw and ready-to-cook poultry products. This method is fast, sensitive, and aligns with FSIS guidelines, enhancing food safety and supporting public health efforts.

Salmonella is a leading cause of foodborne illness in the United States, particularly in poultry products. Traditional methods for detecting Salmonella ...

A Hybrid DNA Extraction Method for the Qualitative and Quantitative Assessment of Bacterial Communities from Poultry Production Samples

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