Urban Microbial Community Dynamics and Metagenomics

Urban Microbial Community Dynamics and Metagenomics

Introduction:

Microorganisms are present in almost every natural environment and, as a group, they are often the most numerous type of organism in an environment. Until recently, studies of microbial diversity were limited to methods where only microorganisms whose growth could be detected in the lab in nutrient media could be identified, quantified and studied. It is estimated that only 5% to 10% of the microbial diversity in any given sample can be cultivated in the lab and detected in this way, and because of this much of the microbial diversity has gone undetected and under appreciated. Recently metagenomic approaches have been introduced that detect a much greater portion of the actual diversity in a sample. Metagenomic approaches are culture-independent, which means that it is not necessary to be able to cultivate the microorganisms in order to detect their presence. Instead, metagenomics requires isolating the genomic DNA (or RNA in the case of some viruses) from the environmental sample being studied and using “signature” DNA sequences as a proxy to quantify and characterize the microorganisms present in the sample. Because these approaches are not selective for specific microbial genomes in a sample the isolated DNA contains genomes representing, in principle, all of the microorganisms that were present in the sample (hence the prefix “meta”, in metagenomics). When a sample is being studied to determine the microbial population diversity and relative abundance in an environmental sample, usually a specific region of each microbial genome is examined – in the case of bacteria this is most often the gene coding for the 16S rRNA. The 16S rRNA gene is used because it is present in all bacteria and it has regions of sequence that are shared among all bacteria as well as variable regions in the gene that differ from species to species.

Metagenomic approaches have now been used to analyze numerous natural environments across the globe as well as the microbiomes of humans and a variety of other organisms. However, urban environments have largely been overlooked. Currently more is known about microbial communities in the arctic and distant regions of the oceans than is known about the microbial diversity in any city – despite the fact that a majority of the world’s population now resides in cities (Yergeau et al., 2010; Tringe et al., 2005). In order to address this lack of knowledge, in this lab we will employ a metagenomic strategy, coupled with next generation sequencing, to analyze the microbial communities present on streets, sidewalks and subway platforms in Brooklyn (in and around the BC campus area). Depending on how the experiments are set-up the metagenomic approach can be used to analyze any microorganisms from the environment, however, for the work in this lab we will begin by focusing our investigation on the bacterial members of the urban microbial community and leave viruses, fungi and other groups microorganisms to future studies. The primary objective is to determine the bacterial diversity and relative species abundance found in common urban environments. A second important objective is to examine how these urban bacterial communities change over time. This is important work as urban microbial communities are likely to serve as a reservoir for a number of human pathogens.

Outline of experimental work:

In order to investigate the urban bacterial communities of Brooklyn you will collect samples from the environment and prepare and analyze these samples in the microbiology lab. Initially you will work with your instructor and the other members of your group to identify the site(s) to be sampled. The samples can be collected using a simple swabbing technique. Each of your samples contains the genomic DNA from all of the bacteria that were isolated during the swabbing process. The genome of each bacterial species is represented proportionally to the number of bacteria of that species that were collected on the swab. For example, a species that makes up 20% of the total bacteria isolated from the environment on the swab will have its genome represent ~20% of the total number of bacterial genomes isolated. After collecting the environmental sample the bacteria need to be released from the swab and the DNA from the bacteria needs to be purified away from the bacterial cells and all the other unwanted dirt and debris that is picked up during the swabbing step. Once the metagenomic DNA sample has been purified it must be quantified. We will use universal PCR primers to amplify a variable region within the bacterial 16S rRNA gene. The universal primers are designed to be variable in several positions (that is, a given position in the sequence of the primer could be A, C, G or T for example, and the resulting primers will not all be identical in sequence). The degenerate primers permit the amplification of 16S rRNA gene from as broad a range as possible of the bacterial species represented in the sample. Following PCR amplification the product can be cleaned up using the QIAquick PCR Purification Kit (Qiagen). Each PCR sample will be visually examined by gel electrophoresis to confirm the correct band size and each sample will be quantified using a NanoDrop 2000 spectrophotometer. This amplified 16S rRNA gene region (referred to as the amplicon) can then be cleaned-up, quantified and sent out for next generation sequencing. The company that sequences our DNA samples uses the Roche/454 DNA pyrosequencing technology (see animation here: http://454.com/products/technology.asp). The sequence data from the 16S rRNA genes can them be compared to a database to determine the bacterial diversity of the sample. These steps are listed here and detailed protocols for each procedure are described below.

1.  Identify sites to test and collect environmental samples

2.  Isolate and purify metagenomic DNA samples

3.  Quantify metagenomic DNA

4.  Amplify a variable region from the 16S rRNA gene from metagenomic sample using universal primers

5.  Analyze and quantify 16S rRNA gene amplicon

6.  Clean-up 16S rRNA amplicon and send out for Roche/454 pyrosequencing.

Required background reading:

N. Pace review article: Mapping the Tree of Life: Progress and Prospects; MICROBIOLOGY AND MOLECULAR BIOLOGY REVIEWS, Dec. 2009, p. 565–576

J. Handelsman review article: Metagenomics: Application of Genomics to

Uncultured Microorganisms; MICROBIOLOGY AND MOLECULAR BIOLOGY REVIEWS, Dec. 2004, p. 669–685

Microbiology; A Human Perspective, 6th edition, Nester et al., chapter 1 and chapter 11 (or equivalent chapters from other introductory microbiology texts).

Recommended* background reading:

Logares, R., Haverkamp, T. H. A., Kumar, S., Lanzén, A., Nederbragt, A. J., Quince, C., & Kauserud, H. (2012). Environmental microbiology through the lens of high-throughput DNA sequencing: Synopsis of current platforms and bioinformatics approaches. Journal of Microbiological Methods, 91(1), 106-113.

Metzker, M. L. (2010). Sequencing technologies the next generation. Nature Reviews Genetics, 11(1), 31-46.

Scholz, M. B., Lo, C. -., & Chain, P. S. G. (2012). Next generation sequencing and bioinformatic bottlenecks: The current state of metagenomic data analysis. Current Opinion in Biotechnology, 23(1), 9-15.

Shokralla, S., Spall, J. L., Gibson, J. F., & Hajibabaei, M. (2012). Next-generation sequencing technologies for environmental DNA research. Molecular Ecology, 21(8), 1794-1805.

* These reading cover next generation sequencing technologies and metagenomics data analysis. They can be technical, but reading them to get the general sense of things can be very useful.

Protocol:

Step 1 and Step 2

1.  Sample collection

a.  Select sites for sampling

b.  Swab from 4 spots within a square meter at the chosen sites

i.  Wear gloves and take precautions to avoid contaminating the site or the swabs from your skin or hair, etc.

ii. Dip a sterile swab into sterile swab solution (0.15M NaCl, 0.1% tween) and then swab an area the size of your palm, or a little larger (~8x8 cm)

iii.  Place the swab in a sterile 15 ml Falcon tube. You may need to break off the end of the swab to fit it in the tube.

iv.  Repeat this with three more swabs in three nearby spots next to where you collected your first sample. Note – collect ALL of your samples from within the 1 m2 space but do not re-swab any area. After swabbing place each swab in the same 15 ml Falcon tube.

c.  Back in the lab - release sampled material from swabs by adding 2 ml of swab solution to the Falcon tube containing the 4 swabs and vortexing for 1 minute.

d.  Remove each swab from the tube, but squeeze out all excess liquid on the side of the tube.

e.  Check the volume remaining in the 15 ml Falcon tube, it will probably be between 0.5 and 1.5 mls remaining in the tube and it should look like dirty water with particulate debris that came along on the swab. Split the volume in the 15 ml Falcon tube equally between two sterile microcentrifuge tubes. Take note of approximately how much volume is in the microcentrifuge tube. Spin for 3 minutes at 14,000 x g (gravity is the same as relative centrifugal force; RCF). This step should pellet all the debris and bacteria (and bacteria bound to debris) into the bottom of your microcentrifuge tube. Make sure to face the hinge of the tube to the outside (away from the center of the rotor – this is so any pellet will collect at the bottom or side of the tube underneath the hinge). While your tubes are spinning, calculate how much supernatant should be removed from your tubes to leave ~100 ul remaining in each microcentrifuge tube.

i.  Remove the calculated amount of supernatant using a pipette and being very careful not to dislodge the pellet. When you are done you should have ~100 ul of liquid (about a tear drop) left in the tube with the pellet of debris and bacteria.

ii. Resuspend the pellet in the remaining ~100 ul of supernatant by briefly vortexing or flicking each of the two tubes.

f.  Follow the Mo Bio Power Soil kit (#12888) protocol for isolation of genomic DNA form your sample. The steps will be listed here, but are essentially identical to those written in the Mo Bio Power Soil protocol.

1.  Transfer the entire resuspended ~100 ul from each of the two microcentrifuge tubes to a single Power Soil PowerBead tube. You are using the combined ~200 ul you collected from the swabs instead of a soil sample. What’s happening: After your sample has been loaded into the PowerBead tube the next step is homogenization and lysis. The PowerBead tube contains a buffer that will (a) help disperse the debris particles, (b) begin to dissolve any humic acids that may be present and (c) protect the nucleic acids (genomic DNA) from degradation.

2.  Gently vortex or invert PowerBead tubes to mix. What’s happening: gentle vortexing mixes the components in the PowerBead tube and begins to disperse the sample in the PowerBead solution.

3.  Obtain solution C1 from your instructor. If solution C1 has a precipitate in it then notify your instructor – it may be necessary to heat the C1 solution to 60 C. What’s happening: Solution C1 contains SDS and other disruption agents required to complete cell lysis of the bacteria in your sample. In addition, SDS is an anionic detergent that breaks down fatty acids and lipids associated with the cell membrane of many bacteria. If it gets cold it will form a white precipitte in the bottom of the tube. If this is the case then heating to 60 C will dissolve the precipitate of SDS and the C1 solution can then be used.

4.  Add 60 ul of solution C1 to your PowerBead tube and invert several times to mix or vortex briefly.

5.  Secure PowerBead tube horizontally using the vortex adapter or, alternatively, use the TissueLyzer. Vortex for 10 minutes at maximum speed (or for Tissuelyzer, shake for 2x 5 minutes at 25 Hz, rotating tube adapter between the two 5 minute shakings). What’s happening: the vortexing step is critical for complete homogenization and cell lysis. Cells are lysed by a combination of chemical agents from steps 1-4 and mechanical shaking. By randomly shaking the beads in the presence of disruption agents, collisions of the beads with the microbial cells will cause the cells to break open.

6.  Place your PowerBead tube in the centrifuge as shown by your instructor. Make sure your tube is balanced against another PowerBead tube (from another group) containing approximately the same volume (if there is not another group’s tube to balance with, then use a microcentrifuge tube containing water that weighs the same as your PowerBead tube). Centrifuge your tube at 10,000 x g for 30 seconds at room temperature. CAUTION: be sure not to exceed 10,000 x g or the tubes may crack.

7.  Transfer the supernatant to a clean, labeled, 2 ml collection tube. Note: expect between 400-500 ul of supernatant at this step. The exact recovered volume depends on the absorbency of your starting material and is not critical for the procedure to be effective. The supernatant may be dark in appearance and still contain some particulate debris. Subsequent steps in the protocol will remove the particulate matter.

8.  Obtain solution C2 from your instructor. Add 250 ul of solution C2 and vortex for 5 seconds. Incubate at 4 C (in the refrigerator) for 5 minutes. What’s happening: solution C2 is a patented inhibitor Removal Technology (IRT). It contains a reagent to precipitate non-DNA organic and inorganic material, cell debris and proteins. It is important to remove contaminating organic and inorganic matter that may reduce the DNA purity and inhibit downstream DNA applications, such as PCR.