20.109(F18):Complete in silico cloning (Day1)

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20.109(F18): Laboratory Fundamentals of Biological Engineering

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Fall 2018 schedule        FYI        Assignments        Homework        Class data        Communication
       1. Measuring genomic instability        2. Modulating metabolism        3. Engineering biomaterials              


Introduction

Though the theme of Module 2 is metabolic manipulation / engineering, today will focus on a few key techniques used in DNA engineering. Because the sequence of proteins is determined by the sequence of the genes that encode them, learning how to manipulate DNA is an important first step. Today you will complete a cloning reaction to generate an expression vector that encodes the catalytically inactive ("dead") dCas9 protein. This process is illustrated in the schematic below. Later you will use this in the CRISPRi system to modulate a mixed-acid fermentation product.

Schematic of pdCas9 cloning. First, the dCas9 insert is PCR amplified to generate multiple copies of the fragment that are flanked by restriction enzymes sites. Next, this fragment and the vector are digested to create compatible ends. Last, the compatible ends of the digested insert and vector are 'glued together' in a ligation reaction.

The vector has several features that make it ideal for cloning and plasmid replication -- both of which are important for this module. To generate your final product you will use three common DNA engineering techniques: PCR amplification, restriction enzyme digestion, and ligation.

PCR amplification

The applications of PCR (polymerase chain reaction) are widespread, from forensics to molecular biology to evolution, but the goal of any PCR is the same: to generate many copies of DNA from a single or a few specific sequence(s) (called the “target” or “template”).

In addition to the target, PCR requires only three components: primers to bind sequence flanking the target, dNTPs to polymerize, and a heat-stable polymerase to carry out the synthesis reaction over and over and over. DNA polymerases require short initating pieces of DNA (or RNA) called primers in order to copy DNA. In PCR amplification, forward and reverse primers that target the non-coding and coding strands of DNA, respectively, are separated by a distance equal to the length of the DNA to be copied. Length is one important design feature. Primers that are too short may lack requisite specificity for the desired sequence, and thus amplify an unrelated sequence. The longer a primer is, the more favorable are its energetics for annealing to the template DNA, due to increased hydrogen bonding. On the other hand, longer primers are more likely to form secondary structures such as hairpins, leading to inefficient template priming. Two other important features are G/C content and placement. Having a G or C base at the end of each primer increases priming efficiency, due to the greater energy of a GC pair compared to an AT pair. The latter decrease the stability of the primer-template complex. Overall G/C content should ideally be 50 +/- 10%, because long stretches of G/C or A/T bases are both difficult to copy. The G/C content also affects the melting temperature. PCR is a three-step process (denature, anneal, extend) and these steps are repeated 20 or more times. After 30 cycles of PCR, there could be as many as a billion copies of the original target sequence.

Kary Mullis.

Based on the numerous applications of PCR, it may seem that the technique has been around forever. In fact it is just over 30 years old. In 1984, Kary Mullis described this technique for amplifying DNA of known or unknown sequence, realizing immediately the significance of his insight.

"Dear Thor!," I exclaimed. I had solved the most annoying problems in DNA chemistry in a single lightening bolt. Abundance and distinction. With two oligonucleotides, DNA polymerase, and the four nucleosidetriphosphates I could make as much of a DNA sequence as I wanted and I could make it on a fragment of a specific size that I could distinguish easily. Somehow, I thought, it had to be an illusion. Otherwise it would change DNA chemistry forever. Otherwise it would make me famous. It was too easy. Someone else would have done it and I would surely have heard of it. We would be doing it all the time. What was I failing to see? "Jennifer, wake up. I've thought of something incredible." --Kary Mullis from his Nobel lecture; December 8, 1993


Restriction enzyme digest

Restriction enzyme digest with EcoRI. EcoRI cuts between the G and the A on each strand of DNA, leaving a single stranded DNA overhang (also called a “sticky end”) when the phosphate backbone is cleaved.

Restriction endonucleases, also called restriction enzymes, 'cut' or 'digest' DNA at specific sequences of bases. The restriction enzymes are named according to the prokaryotic organism from which they were isolated. For example, the restriction endonuclease EcoRI (pronounced “echo-are-one”) was originally isolated from E. coli giving it the “Eco” part of the name. “RI” indicates the particular version on the E. coli strain (RY13) and the fact that it was the first restriction enzyme isolated from this strain.

The sequence of DNA that is bound and cleaved by an endonuclease is called the recognition sequence or restriction site. These sequences are usually four or six base pairs long and palindromic, that is, they read the same 5’ to 3’ on the top and bottom strand of DNA. For example, the recognition sequence for EcoRI is below (see also figure at right). EcoRI cleaves the phosphate backbone of DNA between the G and A of the recognition sequence, which generates overhangs or 'sticky ends' of double-stranded DNA.

5’ GAATTC 3’
3’ CTTAAG 5’

Unlike EcoRI, some other restriction enzymes cut precisely in the middle of the palindromic DNA sequence, thus leaving no overhangs after digestion. The single-stranded overhangs resulting from DNA digestion by enzymes such as EcoRI are called sticky ends, while double-stranded ends resulting from digestion by enzymes such as HaeIII are called blunt ends. HaeIII recognizes

5’ GGCC 3’
3’ CCGG 5’

Ligation

Schematic of DNA ligation.

In a ligation reaction, DNA ends are covalently attached to one another via the ligase enzyme. The efficiency of the reaction is related to type of DNA ends: compatible sticky ends will ligate more efficiently than blunt ends, and non-compatible sticky ends will not be ligated due to the lack of hydrogen bonding between the basepairs. To initiate the ligation reaction, hydrogen bonds are formed between the compatible overhangs of DNA fragments. The ligase enzyme then forms a covalent phosphodiester bond between the 3' hydroxyl end of the 'acceptor' nucleotide and the 5' phosphodiester end of the 'donor' nucleotide.

The first step in this process is the addition of AMP (adenylation) to a lysine residue within the active site of DNA ligase, which releases a pyrophosphate. Next, the AMP is transferred to the 5' phosphate of the donor nucleotide resulting in the formation of a pyrophosphate bond. Lastly, a phosphodiester bond is formed between the 5' phosphate of the donor nucleotide and the 3' hydroxyl of the 3' acceptor nucleotide.


Protocols

Part 1: PCR amplification and restriction enzyme digest of dCas9 insert

Because DNA engineering at the benchtop can take days, if not weeks, you will generate your clone in silico today. You can use any DNA manipulation software you choose to complete the protocols, but the instructions provided are for Benchling, the same program you use for your laboratory notebook. Please note that if you use a different program the teaching faculty may not be able to assist you.

Be sure to document your work and answer all questions in your lab notebook as you progress through the exercises below.

PCR amplification and restriction enzyme digest of dCas9 insert.

To amplify a specific sequence of DNA, you first need to design primers -- one primer that anneals at the start of the sequence of interest and a second primer that anneals at the end of the sequence of interest. Today you will design a 'forward' primer that anneals to the non-coding DNA strand and reads toward the IPC gene and a 'reverse' primer that anneals to the coding DNA strand at the end of the IPC gene and reads back into it. Each primer will consist of two parts: the 'landing sequence' will anneal to the sequence of interest and the 'flap sequence' will be used to add a restriction enzyme recognition sequence to your dCas9 insert.

  1. Find the dCas9 insert sequence here.
    • Open Benchling. On the left panel, click the '+' sign and and choose DNA Sequence--> Input Raw Sequence.
    • Type in "dCas9" for the name, "linear" for the topology, click the appropriate folder, and click "Create."
    • Copy and paste the sequence from the .docx file above.
    • Record the size of the dCas9 gene in your notebook.
  2. Because we want to amplify the entire gene, the landing sequence of the forward primer will begin with the first basepair of the sequence.
    • Record the first 20 basepairs of the dCas9 gene sequence in your notebook.
  3. We will use Benchling to assess the characteristics of your primer:
    • Highlight the primer sequence in benchling, right click, and select "Forward primer"
    • Leave the default settings for now, and click on "Check secondary structure"
    • Use the following guidelines to evaluate your primer:
      • length: 17-28 basepairs
      • GC Content: 40-60%
      • Tm: 60-65°C
      • Check for hairpins, complementation between primers, and repetitive sequences (you can click on "All structures" to look at possible homodimers and hairpins).
    • If you primer does not fit the guidelines provided above, try altering the length. Remember that the 5’ end of the landing sequence must not change or you will delete basepairs from your gene.
    • When you are satisfied with the landing sequence, either save it with the name "landing sequence" or make an annotation labeled "landing squence" according to the following instructions.
      • Highlight the landing sequence you decided on
      • Click Create → Annotation then complete the requested information in the Annotations window.
      • Click Save Annotation.
  4. Now that you have your landing sequence you will add a flap sequence that introduces a restriction enzyme recognition sequence.
    • As shown in the schematic of our cloning strategy, we need to add a BglII recognition sequence to our forward primer. Search the NEB list to find the BglII recognition sequence. Record the recognition sequence and the cleavage sites within the sequence.
    • Add the recognition sequence for the BglII restriction enzyme to the landing sequence. Consider the direction in which PCR amplification occurs to determine which end of your primer should carry the flap sequence.
    • In addition to the recognition sequence, it is important to include a 6 basepair 'tail' or 'junk' sequence to ensure the restriction enzyme is able to bind and cleave the DNA. Learn more about why this is necessary from scientists at NEB. Add any sequence of 6 basepairs to your primer flap sequence. Carefully consider where this sequence should appear in your primer.
  5. Record the sequence (5' → 3') of your forward primer in your notebook.
  6. Use steps 2-5 to design your reverse primer. Please keep the following notes in mind:
    • Because you want to amplify the entire gene you should start with the last basepair of the sequence.
    • You will add an XhoI restriction recognition site to your reverse primer.
    • Remember that the reverse primer anneals to the coding DNA strand at the end of the dCas9 gene and reads back into it. Keep this in mind when you add the flap sequence and when you record the sequence (5' → 3') of your primer in your notebook.
  7. Highlight the entire sequence (of dCas9) and click Copy.
    • In the window that opens, click on the Sequence box.
  8. Paste the sequence into a new Sequence Map window ('+' Create → DNA Sequence → Input Raw Sequence) that depicts the dCas9 product you would expect if you used your primers in a PCR amplification reaction.
    • Save this sequence as 'dCas9 PCR insert'.
    • Be sure to include the restriction enzyme and junk sequence that is added by your primers.
    • What is the size of your PCR product? How does this compare to the size of the gene you recorded in Step #1.
  9. Now that you have your amplified dCas9 insert, you need to digest with BglII and XhoI to generate 'sticky ends' that will enable you to ligate your insert into the vector.
  10. Create another new sequence that depicts your amplified dCas9 PCR insert product following a BglII and XhoI double-digest.
    • What is the size of your digest product? How does this compare to the size of your PCR product?

Part 2: Restriction enzyme digest of vector

To prepare for the ligation step, it is important to generate compatible 'sticky ends' on the insert and vector. Above, you digested your dCas9 amplicon (PCR amplification product) with BglII and XhoI in a double-digest. Here you will digest your vector to create compatible ends that can be ligated together.

Restriction enzyme digest of vector.
  1. Find the vector sequence here.
    • Copy and paste the vector sequence into a new Sequence Map window and save this sequence as 'expression vector'.
    • Be sure to select Circular from the Topology dropdown.
  2. Cloning vectors are engineered to contain a Multiple Cloning Site (MCS). The MCS is a short segment of DNA that encodes several restriction enzyme recognition sites. These restriction enzyme recognition sites are provided for so researchers can clone their genes of interest into a specific location of the vector.
    • Using the Annotation function (ribbon symbol on right side), label basepairs 734 to 783 as the MCS.
    • Label basepairs 722-733 as ribosomal binding site (RBS).
    • Label basepairs 1077 to 1622 as p15A ori. This is the origin of replication.
    • Label basepairs 1851 to 2510 as chloramphenicol acetyltransferase (camR). This gene is provides chloramphenicol resistance.
  3. At the top panel, click Plasmid to see a visual representation of your vector map.
  4. To 'digest' your vector for cloning, Click on the scissor icon at the far right of your screen.
    • A New Digest window will open.
    • Enter BamHI into the Find Enzyme box.
    • Click on BamHI and enter XhoI into the Find Enzyme box. Both BamHI and XhoI (and no other enzyme) should be listed under the Selected header.
    • Click Run Digest.
  5. Of the two fragments generated in your digest, which is the vector backbone that you will use for cloning? Which is the removed MCS?
    • Save your digest by entering 'BamHI-XhoI' in the box at the top of the window then click Save.

Part 3: Ligation of dCas9 insert and expression vector

When you complete a ligation at the bench, one very important step is to calculate the amounts of DNA you will use in the reaction. Ideally, you would use a 4:1 molar ratio of insert to backbone, and would need to calculate how much volume of each solution to use. You can use the steps below to calculate the amount of dCas9 insert and the vector you would use to complete this ligation in the laboratory.

Recovery gel for ligation calculations. Lane 1 = dCas9 insert, Lane 2 = molecular weight ladder, and Lane 3 = cloning vector.
  1. The concentrations for the insert and vector were measured using a nanodrop.
    • dCas9 insert = 40 ng/mL
    • cloning vector = 20 ng/mL
  2. Convert the mass concentration to a molar concentration, using the fact that a typical DNA base is 660 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 math better that way.
    • Hint: you need to know the number of basepairs in the backbone and insert. Use your text sequences and/or Benchling files.
  3. Ideally, you will use 50-100 ng of backbone in the this ligation.
    • Referring to the mass concentration, what volume of DNA will this amount require?
  4. 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?
  5. A 15 μL scale ligation should not include more than 13.5 μL of DNA because you must leave enough volume to add buffer and the ligase enzyme.
    • 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.
  6. Be sure to record all of your work for the ligation calculations in your notebook.
    • Feel free to take a picture of your hand-written work and embed the image in your notebook.
  7. Next you will complete this ligation in silico to generate a plasmid map of your pdCas9 plasmid.
    Ligation of dCas9 insert and cloning vector.
  8. To ligate your dCas9 PCR insert into the expression vector, be sure the vector sequence is in the Sequence Map window.
    • Click the Clock icon on the far right to open the History window.
    • Under the header Clone Version to New DNA enter pdCas9 in the Name box.
    • Select Clone Version.
  9. Open the dCas9 PCR insert in the Sequence Map window.
    • Click the Gear icon at the top of the window and be sure that Cut Sites is checked.
  10. Select the BglII label on the sequence, then hold the Shift key and select the XhoI label. The insert sequence should be highlighted.
    • Click Copy and In the window that opens, click on the Sequence box.
  11. Go back to the pdCas9 sequence and select the BamHI and XhoI labels as in Step #10.
    • Use the keystroke Command + V (for Mac) or Control + V (for PC) to 'ligate' the dCas9 PCR insert into the expression vector, thereby generating pdCas9.
  12. Annotate the dCas9 PCR insert within the ligation product as above.
  13. Use the pdCas9 plasmid map to answer the following questions.
    • What is the size (in bp) of your ligation product?
    • Does your sequence still contain a BamHI recognition sequence? A BglII recognition sequence? Explain.
    • Does your sequence still contain a XhoI recognition sequence? Explain.

Part 4: Confirmation digest

To confirm the pdCas9 construct that we will use for this module, you will perform a 'diagnostic' or 'confirmation' digest. Recall from lecture that this step is important as a control -- you want to be sure that the products you use in your research are correct. This is an important step to check products you clone yourself and, perhaps more importantly, those that you may receive from another researcher.

Ideally you will use a single enzyme that cuts once within the vector and once within your insert. Unfortunately, this is rarely an option and you instead need to select an enzyme that cuts once within the vector and a second, compatible enzyme that cuts once within the insert. Enzyme compatibility is determined by the buffer. If two enzymes are able to function (cleave DNA) in the same buffer, they are compatible. The | NEB double digest online tool will prove very helpful!

Use information from the lecture, the 20.109 list of enzymes and the plasmid map you generated above to choose the enzymes you will use.

  1. To choose restriction enzymes for your confirmation digest, look at the plasmid map for your pdCas9 ligation product.
    • Identify possible sites that will enable to you confirm the pdCas9 sequence.
    • Remember the guidelines discussed in lecture!
  2. After you decide on the enzymes you will use for your confirmation digest, generate a virtual digest in Benchling.
    • Click on the scissor icon at the far right of your screen.
    • Under the Find Enzymes header enter the name of your first enzyme then click on it and enter the name of your second enzyme and click on it.
    • Click Run Digest.
  3. Save the digest as above.
    • What are the sizes of your digest fragments?
    • What buffer should you use for your digest reactions?
  4. Click on the Virtual digest tab at the top of the window to see a 'gel' image.
    • Are the fragments distinct or ambiguously close together?

Keep the following in mind as you consider which enzymes to use:

  • Each enzyme should be present in 2.5 U quantity. As an example, the XbaI vial contains 20,000 U/mL, or 20 U/μL, that is to say 8 times the desired working quantity in one microliter; therefore one reaction will require 0.125 μL.
  • Because the lower limit of your pipet is 0.5 μL, you will need to dilute the enzyme in its appropriate buffer prior to adding it to your master mix.
  • The 20.109 enzyme stocks are always the "S" size and concentration.

The following table may be helpful as you plan your work:

Diagnostic digest Enzyme 1 only Enzyme 2 only No enzyme (uncut)
pdCas9 10 μL 10 μL 10 μL 10 μL
10X NEB buffer 2.5 μL of buffer#_____ 2.5 μL of buffer#_____ 2.5 μL of buffer#_____ 2.5 μL of buffer#_____
1st Enzyme (2.5 U) ____ μL of _____ ____ μL of _____
2nd Enzyme (2.5 U) ____ μL of _____ _____ μL of _____
H2O to a final volume of 25 μL
  1. Unlike the cloning steps you completed above, the diagnostic digest will be performed at the benchtop.
  2. Prepare a reaction cocktail for each of the above reactions (uncut, singly cut with enzyme 1, singly cut with enzyme 2 and doubly cut with enzyme 1 and enzyme 2) that includes (in that order) water, buffer and enzyme.
  3. Aliquot 10 μL of pdCas9 into four well-labeled eppendorf tubes.
    • The labels should include the plasmid name, the enzymes to be added, and your team color.
  4. Add 15 μL of the appropriate cocktail to each tube. Flick the tubes to mix the contents then gather the liquid in the bottom of the tube with a short spin down.
  5. Incubate your digests at 37 °C.

The teaching faculty will leave your digests at 37 °C for one hour, then move them to -20 °C.

Reagents

  • pdCas9 (concentration: 0.05 μg / μL)
  • NEB buffer
    • The buffer will depend on the enzymes you use for your confirmation digest, but all NEB buffers are supplied at a 10X concentration.
  • NEB enzymes
    • The concentrations for each enzyme are listed on the product information page of the website.

Navigation links

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