20.109(S14):16S PCR and paper discussion (Day3)

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#Ligation calculation in advance of Day 4 in progress…
#Ligation calculation in advance of Day 4 in progress…
#Day 4 of this module is poised to run long, so you should read Parts 2 and 3 of the protocol in full and perhaps prepare some of your lab notebook in advance. In addition, to save time later you should prepare an automated worksheet (e.g., in Excel) that will perform the required calculations for that day. Your worksheet should do the following, and a copy must be handed in using the mock numbers provided:
#Day 4 of this module is poised to run long, so you should read Parts 2 and 3 of the protocol in full and perhaps prepare some of your lab notebook in advance. In addition, to save time later you should prepare an automated worksheet (e.g., in Excel) that will perform the required calculations for that day. Your worksheet should do the following, and a copy must be handed in using the mock numbers provided:
-
#*First, prepare the known elements: the vector size is 3500 bp and the concentration is 25 ng/μL.
+
#*First, prepare the known elements: '''the vector size is 3500 bp and the concentration is 25 ng/μL'''.  
-
 
+
#*Another known element, but one that you must calculate yourself based on Day 3, is the size of the PCR product. Scroll down to the reagent list and note that the primers are named according to the base-pair sites that they span.
-
F13 to revise
+
#*Next, prepare spaces for variable elements: desired molar ratio; concentration of insert; volume of both vector and insert; mass of both vector and insert .
-
 
+
#*For the desired molar ratio, begin with 10. For volume of vector, begin with 1 μL, and calculate the mass.
-
#*Separately calculate the concentration of backbone and of insert based on the recovery gel posted on today's Talk page. Refer to the [https://www.neb.com/products/N3232-1-kb-DNA-Ladder NEB marker] definitions to estimate the ''ng'' of DNA in each lane, and refer to your notebook/protocol for the relevant volume basis. Note that the ''ng'' listed are for 10 μL of ladder.
+
#**'''Note: The range of use for the vector is 5-25 ng per reaction.'''
-
#*You may convert the mass concentration to a molar concentration, using the fact that a typical DNA base is 500 g/mol. This conversion will mostly cancel out between the insert and the backbone, except for the difference in number of bases. Feel free to either omit steps that will cancel if you are comfortable doing so, or to keep them if you follow the lab math better that way.
+
#**'''Note: The molar ratio range is 2-10x, with higher generally being better and X being known to work in our system.'''
-
#*Ideally, you will use 50-100 ng of backbone in the upcoming ligation. Referring to the mass concentration, what volume of DNA will this amount require?
+
#*You will determine the insert concentration by the appearance of the gel that you run on Day 4. Familiarize yourself with the band sizes and associated masses of the [https://www.neb.com/products/N3232-1-kb-DNA-Ladder NEB marker] that we will use.
-
#*Ideally, you will use a 4:1 '''''molar''''' ratio of insert to backbone. Referring to the molar concentrations, how much insert do you need per μL of backbone?
+
#*What if your insert looks similar in brightness to the 2 Kbp band? What mass of insert is in that lane?
-
#*A 15 μL scale ligation should not include more than 13.5 μL of DNA. If your backbone and insert volumes total to greater than this amount, you must (1) scale down both DNA amounts, using less than 50 ng backbone and/or (2) stray from the ideal 4:1 molar ratio. You may ask the teaching faculty for advice during class if you are unsure what choice is best, but make and submit an initial ligation plan for now.
+
#** If you are unsure, assume 20 ng.
-
 
+
#*Peeking ahead at the Day 4 protocol, determine what volume of DNA was in the lane, taking into account dilution by the loading dye.
-
S12 style
+
#** If you are unsure, use 10 μL.
-
 
+
#*From the mass and volume of insert, calculation its mass concentration.
-
#*The worksheet should accept 260 nm and 280 nm absorbance readings for RNA samples.  
+
#*Based on the amount of vector and molar ratio listed above, what volume of insert do you want to use in the ligation reaction?  
-
#**Mock numbers: 0.605, 0.312 for sample 1; 0.567, 0.288 for sample 2.
+
#**Perhaps the most efficient route to this calculation is to begin with the ng of vector to be used, and in about 5 steps multiply ratios and cancel units until you end up with a volume of insert.
-
#*The first calculation should be the 260:280 ratio.
+
#**Note that a typical double-stranded DNA base-pair is about 660 ng. If you approach this calculation correctly, this number should overall cancel out in your equation.  
-
#*The next parameter that the user should be able to specify is the RNA dilution factor. Calculate what this is based on the Day 4 protocol.  
+
#*Make sure that your final answer passes a sanity check. How does mass of a piece of DNA versus moles of a piece of DNA scale with size?
-
#**If you cannot figure it out, use a value of 200 for now.
+
#*Finally, note that in your real (rather than hypothetical) reactions, '''you cannot use more than 5 μL of insert'''.
-
#*The next calculation should be the RNA concentration in μg/mL. Find the conversion factor for absorbance to RNA concentration in the Day 4 protocol.
+
==Reagent list==
==Reagent list==

Revision as of 16:14, 13 February 2014

20.109(S14): Laboratory Fundamentals of Biological Engineering

Home        Schedule Spring 2014        Assignments       
Module 1        Module 2        Module 3              

Contents

Introduction

Last week you prepared a DNA pool extracted from a bird cloacal swab, and today you will amplify 16S rDNA amplicons from that pool using PCR. Recall that PCR consists of repeated melting, annealing, and extension steps. During the annealing step of PCR, primers should largely bind to the target sequence, but some may bind to off-target sequences as well. Specificity of binding is controlled by both primer design and reaction conditions.

During the reaction itself, we can improve the reliability and accuracy of PCR by two key methods. The first is the use of a highly specific polymerase, one with either engineered or inherent hot start properties. Hot start means that the polymerase has reduced at low temperatures. Negligible activity at room temperature allows reaction assembly without chilling, while relatively low activity at typical annealing temperatures (50-55 °C) reduces binding of primers to non-target DNA. Eliminating non-specific binding is especially important when the target DNA may be present at a low concentration compared to the DNA in the sample as a whole, as in a complex mixture. Note that only engineered polymerases, such as those bound to accessory antibodies when at room temperature, are officially called "Hotstart." The polymerase you will use has much lower activity at typical annealing temperatures than does the laboratory workhorse Taq, but does so through an inherent rather than external mechanism.

The second performance enhancer we will use is bovine serum albumin (BSA), which is an especially important additive for amplifying DNA originating from a cloacal swab. Recall that these swabs include feces that are replete with inhibitors, including those that bind directly to DNA polymerases. As you saw if you clicked on the Kreader paper linked on Day 1, BSA itself binds many inhibitors of PCR, thus acting as a competitor. In other words, inhibitors should bind the BSA, rather than bind the polymerase and interfere with its function! BSA is hydrophobic and somewhat positively charged, making it a great non-specific binder of proteins that we will use time and again in 20.109.

Next time, you will visualize your entire reaction mixture in a gel, and if need be excise and purify the band of the correct size (~1400 bp) to isolate it from any non-specific products that occur in spite of the above precautions. Gel electrophoresis is a technique used to separate large molecules by size using an applied electrical field and appropriate sieving matrix. DNA fragments are typically separated in gels composed of agarose, a seaweed-derived polymer (see figure, below left). To prepare these gels, molten agarose is poured into a horizontal casting tray containing a comb. Once the agarose has solidified, the comb is removed, leaving wells into which the DNA sample can be loaded. The loaded DNA samples are then pulled through the matrix when a current is applied across it. Specifically, DNA molecules are negatively charged due to their phosphate backbones, and thus travel toward the positive charge at the far end of the gel (see figure, below right).

Scanning EM image of agarose polymer
Scanning EM image of agarose polymer


Although all DNA molecules travel in the same direction during gel electrophoresis, they do so at different rates: larger molecules get entwined in the matrix and retarded, while smaller molecules wind through the matrix more quickly and thus travel further from the well. Ultimately, fragments of similar length accumulate into “bands” in the gel. Bands of DNA are usually visualized by adding the fluorescent dye ethidium bromide (or newer alternatives such as SYBR Safe) to agarose gels. This dye intercalates between the bases of DNA, allowing DNA fragments to be located in the gel under UV light and photographed. The intensity of the band reflects the concentration of molecules that size, although there are upper and lower limits to the sensitivity of dyes. Because of its interaction with DNA, ethidium bromide is a powerful mutagen and will interact with the DNA in your body just as it does with any DNA on a gel. You should always handle all gels and gel equipment with nitrile gloves. Agarose gels with ethidium bromide must be disposed of as hazardous waste.

One parameter that affects the way DNA travels through a gel is the pore size, which is in turn affected by both the weight percent of the gel and the type of agarose used. Because we are separating large DNA fragments (> 1 Kbp) in the bacteria experiment, a low-to-medium percentage gel (namely 1.5 %) is appropriate. In the microsporidia experiment, we expect small fragments (~ 0.1 Kbp) and thus will use a high percentage gel (namely 3%). Moreover, we will use high-resolution (HR) agarose; its low viscosity means that high weight percent solutions are tractable to work with, and that the solidified gel remains pliable rather than brittle. HR agarose can be prepared by chemically modifying and/or partially depolymerizing natural agarose (as described here).

The 16S PCR will continue during the whole lab period. In the meantime, we will discuss a journal article, both to learn more about investigations of the microbiome and to become comfortable reading and discussing the primary scientific literature. You will also hear from our oral presentation instructor on how to give a good talk, and get immediate feedback on your informal presentation of one slide together with your partner. In 2-3 weeks, you will each present an article on your own.

Protocols

Part 1: Prepare PCR to detect bacterial 16S

  1. Begin by carefully labeling each PCR tube that you will use with the date, sample name, and a unique symbol and/or color for quick identification. Filling in the cap tab with your team color will usually suffice.
  2. Pre-chill the tubes on a cold block.
  3. You and your partner can now prepare and share a so-called "master mix," which contains every PCR ingredient except the template and/or primers and/or polymerase. Here, you will omit only the template since we are all using the same primers. The dNTPs must be added before the polymerase. Prepare enough for the number of reactions you need to run, plus an additional 10%. In addition to the two DNA-containing reactions, you should prepare a no-template control that contains pure water without any plasmid. Feel free to use the table below for your calculations.
    • When the master mix is not in use, keep it on ice.
    • What do you expect to see in the no-template control case?
  4. Combine 45 μL of master mix and 5 μL of template in a PCR tube. When everyone's reactions are ready, they will undergo the cycling conditions listed below.
    • Add the master mix first, because the template alone may freeze. Then add template and polymerase, and finally (gently!) mix the reaction with a larger pipet.
Reagent Amount for 1 reaction (μL) Amount for 3 reactions + 10%
PfuUltra buffer (10X stock) 5
Primer mix 1
dNTPs 1
1% BSA (100X stock) 0.5
Water 36.5
PfuUltra polymerase 1
DNA template 5 N/A
Segment Cycles Temperature (° C) Time
1 1 95 5 min
2-4 35 95 1 min
51 1 min
72 2 min
5 1 72 10 min
6 1 4 indefinite

Parts 2 + 3: Journal article discussion and WAC session

Scientific papers are dense and often time-consuming to read and understand, but with practice, you will find strategies that improve your comprehension efficiency. Here's one tip to get you started: when reading newly reported results, be sure to refer to the associated figures frequently, because visual information is often easier to take in than purely verbal descriptions.

Technical Background

Several terms and approaches in the Koenig et al. paper may be unfamiliar to you. Here we will provide some background on selected topics. You should feel free to search for information about additional topics – even Wikipedia can be a good start! At the same time, don't feel the need to understand every detail presented in the paper.

Named for the company that developed the technique, 454-pyrosequencing is one example of a next generation sequencing method. Such methods were designed to enable efficient and accurate identification of many, many sequences at once. You can learn about this particular approach at the company website.

Metagenomics refers to the investigation of microbiome diversity via all genes occuring in an environmental sample – as opposed to via one gene (usually 16S rRNA) alone. You can learn more in the linked review papers here and here if you wish.

Prof. Runstadler will address 16S rRNA approaches in lecture, and you can also refer to the Day 1 and Day 2 wiki introductions.

You can learn about UniFrac software at the linked paper or the direct site. It was used by the authors for some of their analyses.

Discussion Topics

Writing

As you read the paper by Koenig et al., consider not only its scientific content, but also the authors' writing style (perhaps not all on one read!). Sketch out answers to the questions below, right on the paper if you wish. Your answers will not be collected, but you may be called on in discussion to share your ideas.

  • What functional elements does the abstract contain? As a whole, did the abstract make you want to read the paper?
  • Now consider the Introduction section.
    • What is the topic and/or function of each paragraph?
    • How closely does this introduction conform to the suggested three-section structure described in the class scientific writing guidelines? Is there too much, or too little, information or emphasis on any particular topic?
    • What purpose(s) do the citations serve?
  • Now consider the Results section.
    • What purpose do the sub-section titles serve? Which ones do so most effectively?
    • Can you find one or more examples of paragraphs with effective introductory and concluding sentences, according to the description here?
    • How about examples with ineffective opening and closing sentences? How might you improve these?
    • Are there any parts in the Results that you think belong in the Discussion instead, according to the descriptions here and here ?
  • Finally, consider the Discussion section.
    • What is the topic and/or function of each paragraph?
    • Is there too much or too little information or emphasis on any particular topic?
    • Have the authors framed their writing to suggest future studies?
    • What purpose(s) do the citations serve?
Content

You were previously assigned one of the topics below to present to and discuss with the rest of the class. We'll now break to listen to Atissa's talk about giving talks, then give you some time to revise the slide that you prepared, and finally go through the slides and topics one by one. Per slide, we'll first discuss scientific content, and then give you feedback about your slide design and presentation -- briefly and informally.

  • Figure 1 – assigned to Pink Team
    • How is phylogenetic diversity (PD) defined? What general trend was observed? How do the authors interpret various exceptions to the trend?
  • Figure 2 – assigned to Silver Team and White Team together
    • Skim the linked paper about UniFrac software, and refer to it to explain the basic idea of principal coordinates analysis to your peers. What additional information is learned in Fig 2 as compared to Fig 1?
  • Figure 3 – assigned to Yellow Team
    • What changes in bacterial population do the authors observe over time and to what life events do they attribute them? Which timespan population would you expect to vary the most among different infants, and which would you expect to be most similar across different infants?
  • Figure 4 – assigned to Blue Team
    • Briefly, define what C-score and checkerboard analysis measure. How did the authors decide whether these values were higher, lower, or the same as might be expected? What primary conclusion do they draw?
  • Figures S1 + S2 – assigned to Orange Team
    • What general trends in SCFA concentrations and bacterial load did the authors find, with respect to time? How did the authors approach error measurement, and how reliable does each dataset appear to you?
  • Figure 5 (focus on 5C) – assigned to Green Team
    • Make sure you understand (and can explain) how to read a correlation matrix before you begin. How do the authors interpret the correlations they find when looking at bacterial diversity and metabolite concentrations together?
    • If you have time to look at 5A and 5B... do the figures appear to accurately reflect the text?
  • Figure 6A + S4 – assigned to Red Team
    • What is the difference between the approach the authors take in 6A and S4? How do they explain the differences in their findings by said methods?
  • Figure 6B + Table S5 – assigned to Purple Team
    • Describe a few trends in gene abundance that the authors found. How easily can you discern these findings from the figure and table? How might you redesign the figure and/or table for easier use by the reader?

We’ll briefly discuss Figures S3 and Table S6 as a group.

For next time

Still in progress, with my apologies! Should be ready by mid-afternoon Thursday.

Because of the holiday next week, this assignment is long. Be sure to start it earlier than the night before our next lab day.

  1. You will write up the work you do in the bird gut microbiota experiment as an abstract and summary report. To help you pace your work, as well as give you feedback early on, you will be required to draft small portions of the report as homework assignments. For next time, you should prepare the overview schematic of your experimental approach.
  2. The other major assignment you will complete during Module 1 is your primer design memo. While the information is fresh in your mind, turn your M1D2 notes into a table appropriate for submission, and write the associated paragraph explaining your design choices and hypothesis. See the Motivation section of memo assignment for details. Note that you are not yet required to write up the "overall strategy" paragraph.
  3. Post your final primer design on the wiki so these may be ordered in time. Please add the word "FINAL" next to/below your team color when you are satisfied with the design.
  4. Ligation calculation in advance of Day 4 in progress…
  5. Day 4 of this module is poised to run long, so you should read Parts 2 and 3 of the protocol in full and perhaps prepare some of your lab notebook in advance. In addition, to save time later you should prepare an automated worksheet (e.g., in Excel) that will perform the required calculations for that day. Your worksheet should do the following, and a copy must be handed in using the mock numbers provided:
    • First, prepare the known elements: the vector size is 3500 bp and the concentration is 25 ng/μL.
    • Another known element, but one that you must calculate yourself based on Day 3, is the size of the PCR product. Scroll down to the reagent list and note that the primers are named according to the base-pair sites that they span.
    • Next, prepare spaces for variable elements: desired molar ratio; concentration of insert; volume of both vector and insert; mass of both vector and insert .
    • For the desired molar ratio, begin with 10. For volume of vector, begin with 1 μL, and calculate the mass.
      • Note: The range of use for the vector is 5-25 ng per reaction.
      • Note: The molar ratio range is 2-10x, with higher generally being better and X being known to work in our system.
    • You will determine the insert concentration by the appearance of the gel that you run on Day 4. Familiarize yourself with the band sizes and associated masses of the NEB marker that we will use.
    • What if your insert looks similar in brightness to the 2 Kbp band? What mass of insert is in that lane?
      • If you are unsure, assume 20 ng.
    • Peeking ahead at the Day 4 protocol, determine what volume of DNA was in the lane, taking into account dilution by the loading dye.
      • If you are unsure, use 10 μL.
    • From the mass and volume of insert, calculation its mass concentration.
    • Based on the amount of vector and molar ratio listed above, what volume of insert do you want to use in the ligation reaction?
      • Perhaps the most efficient route to this calculation is to begin with the ng of vector to be used, and in about 5 steps multiply ratios and cancel units until you end up with a volume of insert.
      • Note that a typical double-stranded DNA base-pair is about 660 ng. If you approach this calculation correctly, this number should overall cancel out in your equation.
    • Make sure that your final answer passes a sanity check. How does mass of a piece of DNA versus moles of a piece of DNA scale with size?
    • Finally, note that in your real (rather than hypothetical) reactions, you cannot use more than 5 μL of insert.

Reagent list

  • PfuUltra polymerase and buffer from Agilent
  • dNTPs from Promega, original stock at 10 mM of each base
  • Primers
    • intermediate stock concentration is 5 μM for each primer (10 μM total), diluted from individual 100 μM long-term stocks
    • F8-27 sequence: 5' AGAGTTTGATCCTGGCTCAG
    • R1392-1407 sequence: 5' ACGGGCGGTGTGTACA

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