20.109(S20):Complete in silico cloning and induce TDP43 protein expression (Day1)
Introduction
Though the theme of Module 1 is focused on screening for small molecules that bind TDP43, 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 TDP43 protein. This process is illustrated in the schematic below. Later you will use the protein you purify to complete a small molecule microarray.
![](/w/images/thumb/f/fb/Sp20_M1D1_cloning_schematic.png/450px-Sp20_M1D1_cloning_schematic.png)
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.
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 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
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: Laboratory orientation quiz
Complete the orientation quiz with your partner. Though you are working with your partner, each student should record all answers on the provided quiz. If you disagree with your partner on an answer, you should write what you think is the correct answer on your quiz.
Good luck!
Part 2: PCR amplification and restriction enzyme digest of TDP43-RRM12 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 SnapGene. 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.
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 gene that encodes TDP43 and a 'reverse' primer that anneals to the coding DNA strand at the end of the dCas9 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 TDP43 insert.
- Find the TDP43-RRM12 insert sequence here.
- Open SnapGene. From the options, select 'New DNA File...'.
- Copy and paste the sequence from the .docx file above.
- Enter in "TDP43-RRM12" for the File Name (in the lower, right corner), select 'linear' for the topology (in the lower, left corner), then click 'OK'.
- A new window will open with a map of TDP43-RRM12 showing the unique restriction enzyme sites within the sequence.
- In later steps you will generate a map of the TDP43 insert cloned into an expression vector. To make the map more visually useful, create a feature that defines the TDP43-RRM12 insert.
- Click 'Sequence' from the options at the bottom of the window.
- Highlight the entire sequence in the window.
- From the toolbar, select 'Features' → 'Add Feature...'
- In the new window name, type TDP43-RRM12 into the 'Feature:' box.
- Select gene from the dropdown in the 'Type:' box and select the right facing arrowhead (this denotes the directionality of the insert).
- Then click 'OK'.
- Next you will use the sequence information to design primers that will amplify the TDP43-RRM12 insert.
- Because we want to amplify the entire sequence, 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.
- To label the primer sequence, highlight the first 20 basepairs in the TDP43-RRM12 insert sequence, then select 'Primers' --> 'Add Primer...' from the toolbar.
- A new window will open asking which strand should be used to make the primer. Before making your selection consider the direction in which DNA is synthesized and to which strand your primer should anneal such that the TDP43-RRM12 insert is amplified during PCR.
- In the 'Primer:' text box, enter a specific name for your forward primer, then select 'Add Primer to Template'.
- The primer should be indicated on the sequence of the TDP43-RRM12 insert by an arrow facing into the sequence.
- Click 'Primers' from the options at the bottom of the window.
- Use the following guidelines to evaluate your primer:
- length: 17-28 basepairs
- GC Content: 40-60%
- Tm: 60-65 °C
- Check for hairpins and complementation between primers by clicking on the name of your primer, then 'Primers' --> 'Analyze Selected Primer...' from the toolbar. Note: this will automatically open window to the IDT DNA OligoAnalyzer tool.
- If your 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, be sure to update the primer labeled on the TDP43-RRM12 seqeuence.
- 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 BamHI recognition sequence to our forward primer. Search the NEB list to find the BamHI recognition sequence. Record the recognition sequence and the cleavage location within the sequence.
- Add the recognition sequence for the BamHI 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 the 'Primers' window, click on the name of your primer. Then select 'Primers' --> 'Edit Primer...' from the toolbar.
- Add the recognition sequence by typing into the text box at the top of the window that contains the primer 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.
- Record the full sequence (5' → 3') of your forward primer in your notebook.
- Use the above process 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 EcoRI restriction recognition site to your reverse primer.
- Remember that the reverse primer anneals to the coding DNA strand at the end of the TDP43-RRM12 insert 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.
- To generate the PCR amplicon from the TDP43-RRM12 sequence and your primers, select 'Actions' --> 'PCR' from the toolbar.
- A new window will open, in the text boxes at the bottom select your forward primer (Primer 1) and reverse primer (Primer 2). Then click 'PCR'.
- Record the length of the amplicon in your laboratory notebook.
- Is the amplicon double-stranded or single-stranded? Is it a blunt end product or sticky end product?
- Lastly, save the amplicon file with a specific name.
- Now that you have your TDP43-RRM12 PCR amplicon, you need to digest with BamHI and EcoRI to generate 'sticky ends' that will enable you to ligate the TDP43-RRM12 insert into the vector.
- On the map of the TDP43-RRM12 PCR amplicon, select the BamHI recognition site by clicking on the enzyme name. Then hold the shift key and select the XhoI recognition site.
- This should highlight the area between the enzyme recognition sites.
- Click the drop-down arrow next to the 'Copy' icon at the top of the window.
- Select 'Copy Restriction Fragment.'
- Click the drop-down arrow next to the 'New' icon at the top of the window.
- Select 'New DNA File...'.
- Paste the restriction fragment from the previous step in the text box, then click 'OK'.
- A new window will open with the digested TDP43-RRM12 insert.
- Record the length of the insert in your laboratory notebook. How does the length of the insert compare to the length of the PCR amplicon.
- Is the insert double-stranded or single-stranded? Is it a blunt end product or sticky end product?
- Save the insert file.
Part 3: 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 to create the insert for your cloning. Here you will digest your vector to create compatible ends that can be ligated together.
- Find the vector sequence here.
- Copy and paste the vector sequence into a New DNA File window and save this sequence.
- Be sure to select circular from the topology options.
- As above, a new window will open listing the features recognized by the SnapGene software.
- Include a list and brief description of the features.
- Select 'Add Features'.
- A new window will open with a map of the vector showing the unique restriction enzyme sites and annotated features within the sequence.
- One feature not recognized is the 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.
- To add the MCS, click 'Sequence' from the options at the bottom of the window.
- Highlight basepairs 734 to 783, then select 'Features' --> 'Add Feature...' from the toolbar.
- In the 'Feature:' text box, enter "MCS".
- To generate the sticky ends that will enable you to ligate the dCas9 insert into the vector, view the map of your vector sequence.
- Select the BamHI recognition site by clicking on the enzyme name, then hold the shift key and select the XhoI recognition site.
- Select 'Actions' --> 'Restriction and Insertion Cloning' --> 'Delete Restriction Fragment...' from the toolbar.
- What is the length of the digested vector product? How many basepairs were removed (compare to the intact cloning vector)?
Part 4: 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 3:1 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.
- The concentrations for the insert and vector were measured using a nanodrop.
- dCas9 insert = 25 ng/uL
- cloning vector = 50 ng/uL
- 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 snap gene files.
- 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?
- Ideally, you will use a 3:1 molar ratio of insert to backbone.
- Referring to the molar concentrations, how much insert do you need per μL of backbone?
- 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 3:1 molar ratio. You may ask the teaching faculty for advice during class if you are unsure what choice is best.
- 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.
- Next you will complete this ligation in silico to generate a plasmid map of your pdCas9 plasmid.
- To ligate you dCas9 insert into the expression vector, select 'Actions' --> 'Restriction and Insertion Cloning' --> 'Insert Fragment...'.
- A new window will open. In the bottom workspace of the window, a cloning schematic will appear showing a vector and insert icon.
- Click on the 'Vector' label. Then in the workspace at the the right of the window, select the vector file from the 'Vector:' drop-down.
- Select the restriction enzymes used to digest the expression vector from the drop-down boxes next to the text boxes that contain 'cut'.
- Next, click on the 'Insert' label at the bottom of the window and complete the steps as done for the expression vector.
- For the insert, use the PCR amplicon file.
- Click 'Clone'.
- A new window will open with the cloned final pdCas9 (the 'p' is for plasmid and 'dCas9' refers to the gene that was cloned) product!
- What is the size of the plasmid? Does this make sense given the lengths of the insert and vector?
- Does your sequence still contain a BamHI recognition sequence? A BglII recognition sequence? Explain.
- Does your sequence still contain a XhoI recognition sequence? Explain.
- Bonus question: Why was the vector not digested with BglII for cloning? (Hint: look at the features on the undigested expression vector map)
Part 5: 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.
- 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!
- After you decide on the enzymes you will use for your confirmation digest, generate a virtual digest in SnapGene.
- On the map of pdCas9, select the first recognition site by clicking on the enzyme name. Then hold the shift key and select the second recognition site.
- Select 'Tools' --> 'Simulate Agarose Gel' from the toolbar.
- Record the expected fragment sizes from the digest in your laboratory notebook.
- 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 10 U quantity per reaction. As an example, the XbaI vial contains 20,000 U/mL, or 20 U/μL.
- The 20.109 enzyme stocks are always the "S" size and concentration when you search for them on the NEB website.
- Because the lower limit of your P20 pipet is 2.0 μL, you may need to use the P2 at the front bench for smaller volumes.
- Enzyme volume should not exceed 10% of the total reaction volume to prevent star activity due to excess glycerol.
- To find the concentration of the enzyme(s) you choose, search the NEB site.
The following table may be helpful as you plan your work:
Diagnostic digest (enzyme #1 AND enzyme #2) |
Enzyme #1 ONLY | Enzyme #2 ONLY | Uncut (NO enzyme) | |
---|---|---|---|---|
pdCas9 | 5 μL | 5 μL | 5 μL | 5 μL |
10X NEB buffer (buffer name:____________) |
2.5 μL | 2.5 μL | 2.5 μL | 2.5 μL |
Enzyme #1 (enzyme name:____________) |
____ μL | ____ μL | ||
Enzyme #2 (enzyme name:____________) |
____ μL | ____ μL | ||
H2O | to a final volume of 25 μL |
- Unlike the cloning steps you completed above, the diagnostic digest will be performed at the benchtop.
- Prepare a mix for each of the above reactions (uncut, cut ONLY with enzyme #1, cut ONLY with enzyme #2, and cut with enzyme #1 AND enzyme #2) that includes (in that order) water, buffer, and enzyme in well-labeled eppendorf tubes.
- The labels should include the plasmid name, the enzyme(s), and your team color.
- Aliquot 5 μL of pdCas9 into the four well-labeled eppendorf tubes.
- Flick the tubes to mix the contents then gather the liquid in the bottom of the tube with a short spin in the microcentrifuge.
- 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 list
- pdCas9 (concentration: 25 ng / μL) (a gift from the Prather Laboratory)
- 10X buffer; the buffer will depend on the enzymes you use for your confirmation digest (from NEB)
- restriction enzyme(s); the concentration of each enzyme is listed on the product information page (from NEB)
Next day: [[]]