Ligate Sticky Ends via DNA Ligation

DNA Ligation of Sticky Ends

Ligation of Sticky Ends, Summary of DNA ligation

We have already discussed a high level view of gene cloning in our Molecular Cloning Guide blog post. However, in that blog post we didn’t delve very deep into how we can perform each of the individual steps. Today’s blog post is about ligation. Ligation is the process by which two pieces of DNA can be glued together to form one piece. So, to begin, let’s assume you’ve already decided on a gene product that you want to clone. You’ve also designed primers and completed PCR on the open reading frame in your donor DNA (this could be genomic or non genomic DNA). Your next steps are to digest the PCR product with restriction enzymes and generate sticky ends. You’ll also want to digest your “shuttle” plasmid to generate complimentary sticky ends which will allow your “insert” DNA to click into position into your vector. It’s like a puzzle piece!

Note: It might be useful to look at our RNA Extraction & Isolation guide if you’re planning on making cDNA related to your gene

The above summary is demonstrated here:
Ligate Sticky ends using Ligase

Sticky Ends Insert into a Shuttle Vector

Only some Restriction Enzymes Create Sticky Ends

As you can figure out, generating sticky ends and complimentary ends is extremely important to the above process. However, several different restriction enzymes are available and each of them has different locations where they cut. Also, the type of cuts that they introduce may be “sticky” or “blunt”. Depending on the cloning strategy you are using, you may mix and match different enzymes to achieve different end goals. Ligation of “sticky ends” is much more efficient than ligation of “blunt” ends. Typically 10-100 times more T4 Ligase is required for blunt ends.

Here’s an image with various restriction enzymes and the kinds of ends they produce. Depending on the type of ends, your DNA ligation will proceed very differently!

Restriction Enzymes for DNA Ligation

Ligate DNA via DNA Ligase

Once the restriction enzyme digestion is complete, you can proceed to the ligation step. But, before you digest anything, make sure you’ve planned everything properly! You need to make sure that the insert will be ligated in the proper direction in the shuttle vector. Only once you’ve vetted your overall strategy, should you proceed to ligation and transformation, etc.

There are several kinds of ligase enzymes but the enzyme produced by T4 bacteriophage-infected E. Coli is the most common one. This ligase is called T4 ligase. Whereas normal E. Coli produce DNA ligase that uses NADH as a cofactor, T4 infected E.Coli produce a ligase that uses ATP as a cofactor. This enzyme will find the 3′ Hydroxyl and 5′ Phosphate within your sticky ends and it will form a phosphodiester linkage. If this is confusing, check out the Polymerase chain reaction (PCR) guide for images on what DNA looks like. This is shown here:

Ligation Protocol for T4 Ligase

Phosphodiester Bond Formation during Ligation

Protocol for Ligation of Transgene Insert into Shuttle Vector

Ligation enables fragments of DNA to be combined, such as the cut ends of transgene inserts and plasmids during cloning. This protocol describes the directional cloning of a XbaI/SalI-digested transgene into a shuttle vector, pAdtrackCMV, via cohesive end ligation.

Materials for DNA Ligation

XbaI/SalI digested, gel-purified insert (approx. 1 kb) and pAdTrack-CMV shuttle vector (approx. 9.3 kb; Plasmid #16405, Addgene)
Quick Ligation Kit (contains DNA ligase and 2X Reaction Buffer; #M2200S, New England Biolabs)
Agarose plate containing ethidium bromide
DNA standards

Ligation Methodology

  1. Estimate the DNA concentration of purified insert and vector preparations by applying 1 µl to an agarose gel plate (+ethidium bromide) alongside a range of DNA standards and visualizing under UV light.
  2. Prepare the ligation mix as follows:

    XbaI/SalI digested pAdtrackCMV 50 ng

    XbaI/SalI digested insert 17 ng

    Add water up to 10 µl total volume.
  3. Add 10 µl of 2X Reaction Buffer and mix.
  4. Add 1 µl of DNA ligase and mix.
  5. Microcentrifuge briefly to settle liquid to the bottom of the tube and incubate at 25°C for 5 min.
  6. Place on ice* and transform into desired bacterial strain.

Tips and Tricks for DNA Ligation

  • This reaction setup is using a digested insert to vector DNA molar ratio of 3:1. Inserts of different sizes will require a different amount to be added. Important ligation control reactions to include are (1) digested vector only and (2) digested insert only.
  • Ligation reactions can be stored at -20°C for future use

Applications of Ligation on SciGine

Construction of PB42 Vectors Via Ligation
Plasmid Construction via PCR and Ligation
Plasmid Ligation and Transformation in Yeast
Construct with Human p275UTR
Different DNA ligation methods discussed

Video Tutorial About Sticky Ends and Ligation

References

He et al Proc Natl Acad Sci U S A. 1998 Mar 3. 95(5):2509-14.
Sticky Ends Explained Well
DNA Ligation Theory
Gaastra et al. Ligation of DNA via T4 Ligase
Tsuge et al. One Step Assembly of DNA fragments

Gene Cloning using Plasmids: Molecular Cloning Intro

Gene Cloning using Plasmids via Molecular Cloning techniques

Gene Cloning with Plasmids: Summary

We all know that DNA is the basic building block of biology. So, how can we make use of DNA to change cell biology? Well, today’s blog post will focus on “gene cloning” — making plasmids (circular DNA strands) so that we can introduce them into bacteria using our previous bacterial transformation method. With a plasmid inside the bacteria, you can a) use bacteria to make copies of the plasmid, b) make new proteins with the transformed bacteria and c) do the same inside mammalian cells using the Calcium Phosphate transient transfection method that we developed earlier. With molecular cloning techniques, we can control biology and make cells do some really cool stuff! Note: this is an overview post and does not have a step-by-step protocol associated with it. I’ll tease apart the different steps in future blog posts.

Molecular Cloning of Plasmids: Primer Design

“Cloning” refers to the process of making a copy of a gene so that we can modify it and see what happens. Remember, if you modify genes, your cells start producing new proteins; these proteins could be therapeutic and/or give your cells some new skills. To start, you’ll probably want to review the PCR protocol & guide to remember how PCR works. Now, let’s say we have a gene that we want to clone already available. The next most important part of PCR based gene cloning is the primer…so to design a primer, we need the following:

  • Hybridization sequence: A series of bases that compliment the bases right before your “target gene” or gene of interest.
  • Leader sequence: A few extra bases for our restriction enzymes to make efficient cuts that don’t overlap with our gene of interest.
  • Restriction sites: Places that we will cut so that we can make the plasmid circular.

Take a look at this image to understand the above plasmid design:
Making primers for Gene Cloning PCR

Molecular cloning primer design

Gene cloning product

Be Careful Designing Plasmid Primers for Gene Cloning

Based on the above image, you can tell that if an enzyme’s restriction site is inside your gene of interest, you cannot use that restriction enzyme because you’ll cut your gene. Also, you’ll be putting this gene into a new plasmid. Make sure that the restriction enzyme you use is compatible with the “multiple cloning site” within this new plasmid. If you end up inserting this gene in random locations, the probability that this plasmid will be incorporated into the bacteria or expanded will be significantly decreased.

Look at the image below to understand these tips:
Gene cloning failure - wrong restriction enzyme

Primer mismatch -- Gene cloning error

Gene cloning with PCR

With the primer already designed, we are ready to clone our gene. The rest of the steps in the gene cloning process are:

  • PCR everything
  • Use restriction enzymes to digest the PCR product
  • Use Gel Electrophoresis to purify the insert and the “vector” (recipient plasmid)
  • Ligate the plasmid
  • Transform bacterial cells
  • Isolate our plasmid for future use
  • Analyze the PCR products

Since we already know how to do PCR from our previous blog post, let’s focus on the other stuff. The first step listed is to digest the PCR product. For this, we will use restriction enzymes and incubate them with the PCR products. If everything was designed properly, we would know exactly where the restriction enzymes will cut the DNA in both the “vector” and the “insert”. Next, we will run these restriction digests on a gel and pick out the bands corresponding to our vector and insert (which we already know the size of). Any other “junk” PCR products will be removed in this step. The vector and insert DNA will then be “ligated” to form our new plasmid. To confirm our gene is in this plasmid, we will transform some bacteria with it on a petri dish. Try to make dilutions of your bacteria so that you can grow colonies of bacteria and pick out colonies later on. With the colonies that you pick out, you’ll want to isolate their DNA and digest it to see if your vector and insert are inside. We’ve already isolated the vector and insert in the past, so it’s simple to find out if our insert is inside the bacteria. Finally, as another confirmation, we will sequence the DNA from the bacteria and confirm that everything exists. We will write more about each of these steps in the future, but we wanted you to see them together, as an overview, in this blog post.

Take a look at the steps below:

Preparing plasmid vectors for Gene Cloning

Electrophoresis and Ligation of Genes using Restriction Digests

Transformation of Bacteria and Isolation of Final Gene Clones

Tips and Tricks with this methodology

PCR

: Make sure you choose the melting temperature to match the part of the primer that binds the “open reading frame” (your gene of interest). If you choose the wrong melting temperature, you might get the wrong PCR products because either a) your Tm was too low and you didn’t split the ORF or b) your temp was too high and you got lots of non specific binding.

DNA Digestion

: Make sure DNA digestion occurs for a long time, preferable overnight, to make sure all your vector and insert products were cut and maximize your ligation in the next step. You may need to use alkaline phosphatase in this step. I’ll speak more about that in the future.

Gel electrophoresis

: During gel electrophoresis make sure that you run the correct controls and *know* what wells relate to each of the digested products. Also, make sure you skip lanes to make cutting the wells easier. After this, you’ll need to quantify your DNA so you have enough for the ligation step. You can use a UV spectrometer for this step.

Ligation

: Ligation also requires you to have several controls. For example, you need a ligation reaction without any insert. This will tell you how much background self-ligation your recipient plasmid has. You also need a ligation with some of the other bands you see during your gel electrophoresis. This will tell you how much contaminant DNA there was in your ligation.

Methods related to Gene Cloning on SciGine

Video about Gene Cloning with Plasmids

Notes from our audience

  • “TA cloning is another approach if cloning doesn’t work in systematic way” –Swapnil Oke on Linked In
  • “I think for completeness I think it would be valuable to also mention a few other plasmid features that are important. I didn’t see mention of ribosomal binding sites (RBS) or origin of replication, etc” – Michael Kim on Linked In. — I plan on write about more details regarding plasmid design and purification in the future. For now, please don’t use the above blog post as a comprehensive guide…more like an overview 🙂

References

Molecular Cloning book about PCR based cloning
Addgene Plasmid Reference, a comprehensive guide
Chaokun et al., Fast Cloning

Bacterial Transformation Protocol with Competent Cells

Bacterial Transformation with Competent Cells

Bacterial Transformation using Competent Cells: Summary

Since we have already learned Calcium Phosphate Transfection with mammalian cells, let’s now focus on bacterial transformation of DNA with competent cells. In general, bacterial cells take up naked DNA molecules or plasmids via a process called transformation. Usually, this happens at a slow rate, but when bacterial cells die in close proximity to others, or when they are stressed, the transformation process occurs at a much higher rate. However, not all bacterial cells can be transformed, so biologists use ‘Competent Cells’ which are more inclined to take up DNA. The end goal of transformation is to get bacteria that have your genes of interest so that they will replicate your genes along with their own. If the bacteria contain your genes of interest, you can use them to mass produce proteins, or just store them for extended periods of time because bacteria are so hardy. A good way to test whether your genes of interest were transformed is to include antibiotic resistance in your plasmid. This way, you can be fairly certain that if your bacteria are resistant to antibiotics, they are also carrying genes of interest to you.

Take a look at how natural transformation works:
Transformation Protocol with DNA

Transformation Biology in Bacteria

For bacteria, survival is key and transformation is one of their survival mechanisms. As biologists, we can make use of this survival mechanism for our benefit as well. To do this, we first incubate our competent bacteria with our plasmid and calcium chloride. Bacterial membranes are permeable to chloride ions, but not to calcium. So, as chloride ions enter the cell, the bacteria tend to swell (because they also intake water with chloride ions). Then we heat the bacteria in a process called ‘heat shock’ such that they turn on their survival genes. This causes the bacteria to uptake the surrounding plasmids. With the right design, this plasmid will then be recognized by bacterial DNA polymerases (remember our PCR Guide ?) and it will be expressed/replicated along with the bacteria’s normal DNA.

Take a look below to understand how biologists transform cells:

What is transformation

Transformation Biology

Selecting Transformed Bacteria with Antibiotic resistance in a plasmid

Selecting for Transformed Bacteria with the Lac Z Operon

Once your target plasmid is inside the bacteria, you still need to separate transformed cells from those that are not transformed. Another key challenge is that the transformation process may lead to some DNA being recombined so that your gene of interest is no longer functional. How do you select for cells that only contain functional target DNA that hasn’t been recombined? The trick is to use both antibiotic resistance and a Lac Z operon. By cloning your plasmid along with a Lac Z operon, you give your cells the ability to make a galactosidase protein. If cells have the galactosidase and you feed them X-Gal, they turn blue; cells without this operon are white. So, you first transform all your cells. Then you feed them IPTG to activate the Lac Z operon and cause cells to produce the galactosidase. Then you add in X-Gal and just pick out the bacteria that have functional Lac Z because the useful cells will be a bright blue!

Check out the figure below:

Transformation in Bacteria with LacZ

Transformation leads to Competent cells with LacZ operon

Bacterial Transformation Protocol

Transformation describes the uptake and incorporation of plasmid DNA into bacteria. Antibiotic resistance genes carried on plasmids allow selection of transformants. This protocol describes the transformation of DH5α E. coli with pAdtrackCMV (a vector carrying kanamycin resistance).

Materials for Bacterial Transformation

Ligation mix (20 µl) – insert ligated into pAdTrack-CMV shuttle vector (Plasmid #16405, Addgene)
DH5α competent cells (includes pUC19 DNA control; #18265017, ThermoFisher Scientific)
LB broth (#10855-021, ThermoFisher Scientific)
LB Agar selective plates (prepare from #22700025, ThermoFisher Scientific) with 50 µg/ml kanamycin (#15160054, ThermoFisher Scientific)

Step by Step Transformation Protocol

  1. Thaw competent cells on ice. Aliquot 50 µl into cooled Eppendorf tubes for each transformation reaction.*
  2. Add 5 µl of ligation mix to each tube.*
  3. Incubate on ice for 30 min.
  4. Heat-shock the cells for 20 sec in a 42°C waterbath.
  5. Place on ice for 2 min.
  6. Add 950 µl of warm LB broth per tube.
  7. Allow cells to recover at 37°C for 1 hour with gentle shaking.
  8. Spread 200 µl and 20 µl of each transformation mix onto warm selective plates.*
  9. Incubate plates overnight at 37°C.
  10.  

Notes on this methodology

  • We will talk about “Ligation” in another future blog post
  • Step 1. Unused cells can be refrozen and stored at -80°C for future use.
  • Step 2. As a transformation control, add 1 µl of pUC19 plasmid to one aliquot of cells (pUC19 confers resistance to ampicillin so will need to be seeded onto different selective plates).
  • Step 8. Transformation mix can be stored at 4°C and plated the next day if required.

Bacterial Transformation Video Tutorial

Applications of DNA Transformation on Scigine

References

Excellent Book about Bacterial Transformation
Guide to Common terms in Transformation – Oklahoma University
Compilation of History of Transformation and related protocols
He et al Proc Natl Acad Sci U S A. 1998 Mar 3. 95(5):2509-14.

Calcium Phosphate Transient Transfection Protocol & Guide

Calcium phosphate Transient Transfection Protocol and Guide

Transfection with Calcium Phosphate: General Summary

Molecular biology tools allow us to understand and manipulate DNA/RNA so that we can change how cells behave. In this next series of posts, let’s learn how to manipulate cells and make them do our bidding. Among the list of methods to learn, the first tool to understand is transfection – the process by which we introduce new DNA into a cell so that we can change what proteins it creates. Specifically, we will focus on Calcium Phosphate transient transfection because it is a common and powerful technique. We can then combine transfection with some of our protein-manipulation techniques to change cell behavior and confirm that our changes actually had an effect: Immunoprecipitation (IP) and Western Blotting. Note that other techniques for transfection including electroporation, DEAE:Dextran based transfection, and lipid mediated transfection will be discussed in the future.

Transient vs. Stable Transfection

When you introduce DNA into a cell, it is possible for the cell to keep the DNA temporarily or permanently. Temporarily, a cell might keep your DNA as a packaged plasmid and express it until it divides. Permanent transfection takes place when the new DNA is integrated into the genome of the cell and it passes the DNA down through cell division into its progeny. It’s difficult to determine when genes will be integrated into the genome (stable transfection) and when they will be kept temporarily (transient transition). There is a lot of luck involved. However, it is possible to only keep cells that have your DNA by selection. Take a look at the image below:

Calcium phosphate Mammalian Transfection

DNA Transfection guide

Transient Transfection vs Stable transfection

Calcium Phosphate Transient Transfection

To introduce DNA into eukaryotic cells such as mammalian cells, we need to neutralize the charge on the DNA. This is because cell surfaces are negatively charged and DNA that is unshielded will be repelled from the cell surface. Even if some DNA enters a cell, the nuclear envelope will also reject the DNA due to its charge. (For a picture of the DNA polymer look at our PCR protocol) So, the classical technique for neutralization of DNA’s charge is to use Calcium Phosphate. The steps for transfection with Calcium Phophate are very straight forward:

  • Generate DNA strand (circular DNA is much easier to introduce)
  • Mix calcium phosphate with DNA and generate nanoparticulate precipitates
  • Incubate with cells
  • Select cells expressing the DNA of interest

Cells will tend to phagocytose the calcium phosphate nanoparticles and, with luck, they will allow the nanoparticles to enter the nucleus. Calcium phosphate transfection works well because of the stability provided by divalent calcium ions. Other methods such as lipofectamine and polyethylene imine based transfection also work similarly by neutralizing DNA’s charge. But lipids offer the additional benefit of making the DNA complex more hydrophobic and hence make it easier for it to pass through the phospholipid bilayer.

The general technique is shown below:

Calcium phosphate Nanoparticles and Aggregates

Phagocytosis of Calcium Phosphate leads to Transfection

Selection Media Confirms Stable Transfection with Calcium Phosphate Protocol

Tips and Tricks when optimizing your Calcium Phosphate Transient Transfection Protocol

Calcium Phosphate based transfection is a standard and well known technique. Calcium divalent ions bind the DNA polymers and neutralize their negatively charged phosphate backbones. However, optimization is necessary to maximize the DNA that is phagocytosed into your cell of choice. The variables that affect this technique’s efficacy are:

  • The pH of the solution: Even differences of 0.1 units will have drastic effects on the efficacy of your transfection protocol.
  • Amount of DNA in the precipitate:Some cell types require a lot of DNA in the precipitate such as primary human foreskin fibroblasts. Others will tend to die instead of uptaking DNA, if you add too much DNA.
  • Incubation time with the precipitate:HeLa and 3T3 cells are relatively easy to transfect within 16 hours. These cells can tolerate DNA nanoparticles for extended periods of time. However, other cell types may need shorter incubation times and may tend to apoptose if exposed too long.
  • Additional glycerol or DMSO shock: It may be useful to “shock” cells with a 10% Glycerol solution or a 10-20% DMSO solution for a short time (~3 minutes). Carefully optimize this time for your particular cell type.
  • Rate of Formation of DNA nanoparticles: High efficiency transfection techniques have been discovered whereby buffers like BBS allow DNA nanoparticles to form slowly and precipitate onto cells. When this happens, cells tend to phagocytose more of the adducts and tend towards higher viability/less toxicity.

To make sure that your DNA is being incorporated into cells, you should include a reporter plasmid such as one with neomycin resistance (neo). You will need to optimize the ratio of neo reporter DNA vs. the DNA you want to include.

Calcium Phosphate Transient Transfection Protocol

Materials for Calcium Phosphate Transfection
HeLa cells
Complete DMEM
DNA (10 – 50 ug per transfection)
2.5 M CaCl2 (#C3306 Sigma Aldrich)
2x Hepes Buffered Saline (0.28 M NaCl, 0.05 M HEPES [#H3375 Sigma Aldrich], 1.5 mM Na2HPO4, pH 7.05 exactly)
PBS
Culture Dish

Materials for BBS Calcium Phosphate Transfection
HeLa cells
Complete DMEM
DNA
TE buffer, pH 7.4 (10 mM Tris-Cl, 1 mM EDTA)
2.5 M CaCl2
2x BES-Buffer (BBS) (50 mM BES [#B9879 Sigma Aldrich], 280 mM NaCl, 1.5 mM Na2HPO4 pH 6.95 exactly)
PBS
Selection Medium

Transfection Protocol Steps

  1. Split cells such that there is space between cells.
  2. Clean DNA by adding in 100% ethanol for precipitation
  3. Dry DNA after aspirating supernatant from ethanol. Use air to make sure it is completely dry.
  4. Resuspend pellet in 450 ul of water with 50 ul of 2.5 M CaCl2 buffer
  5. Put 500 ul of 2x Hepes Buffered Saline in a 15 ml conical falcon tube
  6. Add the DNA/CaCl2 solution dropwise to this tube while agitating with a stir bar or other mechanism.
  7. Allow the precipitate to sit at room temperature for 20 minutes
  8. Spread the precipitate over the cells along with their medium . Shake gently to make sure the precipitate is even.
  9. Incubate in cell culture incubator at 37 oC with 5% CO2 for up to 16 hours
  10. Remove medium, wash twice with PBS, and feed cells with complete medium.
  11. Plate cells in selective medium.

BBS High Efficiency Transfection Protocol Steps

  1. Seed cells in a dish so that they can double atleast twice so they can be stably transfected
  2. Next day, dilute DNA in TE Buffer at 1 ug/ul
  3. Make a 0.25 M CaCl2 stock
  4. Mix 20-30 ug of DNA with 500 ul of 0.25 M CaCl2 stock.
  5. Add 500 ul of BBS to this mixture and vortex. Incubate at room temp for 20 min.
  6. Add this mixture to the cell culture dish dropwise and mix by gently shaking the plate.
  7. Incubate cells overnight for 24 h at 3% CO2 at 35 oC.
  8. Wash cells 2x with PBS and then incubate them in complete medium for 2 doublings.
  9. Split cells and incubate in selection media.

Notes on this transfection methodology

  • Cell density has to be low but not too low. The ideal cell density allows you to reach confluence at the end of the transfection period without making the media acidic.
  • Also, space between cells increases transfection efficiency because DNA phagocytosis is proportional to exposed surface area of cells.
  • For some cells, incubate with 10% glycerol or 10-20% DMSO for 3 minutes, and wash twice with PBS, prior to adding the DNA nanoparticles. This may increase your transfection efficiency. However, for the BBS method, this step is not necessary because it will not affect cell transfection efficiency.
  • Supercoiled DNA and plasmid DNA works best with these procedures.
  • Depending on the plasmids that you introduce into cells, your selective medium will vary.
  • pH is EXTREMELY CRITICAL for transfection procedures. At the end of the transfection, pH of your medium should be alkaline at 7.6, but prior to the procedure, make sure all your buffers are clean and at the right pH.
  • All buffers above may be frozen and stored as aliquots. But it is important to make sure that your pH is correct prior to using freshly thawed buffers.

Applications of Transfection on SciGine

Murine L cells transfected with Calcium Phosphate and BBS Buffer
HeLa cell transfection with Lipopolyamine
Calcium Phosphate with HEPG2 and HEK293
MDAMB436 cells with Calcium Phosphate Transient Transfection
Transient Transfection Protocol for HEK293T cells

Transient and Stable Transfection Video Tutorial

References

Calcium Phosphate Transfection by Kingston et al.
High Efficiency Transfection by Chen et al.
Transfection Review by Kim et al.

RNA Extraction, Isolation, and Purification by SciGine

RNA extraction, purification, and isolation

RNA Extraction: General Summary

RNA extraction and isolation is a precursor for many methods in molecular biology including Northern Blotting, RT-PCR, and Microarray analysis. This blog post will focus on this precursor method as opposed to other biology techniques on the Scigine blog where I’ve focused on direct analytical techniques. Nonetheless, there is a lot to learn about RNA isolation and plenty of theory that might be applicable in your research.

Important aspects related to RNA Extraction

While everyone knows DNA is double stranded and helical, few people know that RNA is typically a single stranded polymer. However, secondary structures do emerge within RNA due to complementary base pairing and structures such as tRNA can be double stranded. Also, due to secondary structures, flexible regions of RNA can actually catalyze the cleavage of phosphodiester linkages in nearby RNA chains.

The base polymer looks like this:
Structure of RNA and Effects on Extraction

The RNA polymer has several ionic groups, inter-chain hydrogen bonds, and it is extremely hydrophilic. All of these forces need to be overcome in order to purify RNA from DNA, carbohydrates, and lipids which have similar structures and physical properties. Polymers of RNA can be short or long, but generally smaller strands that are less than 100 nu cleotide bases are hard to purify because they don’t separate well. Additionally, there are several RNAses present in cells and tissues that can catalyze cleavage of RNA chains, so extreme care needs to be taken to prevent degradation of RNA during RNA isolation and purification procedures.

RNA Extraction Method Guide

Typically RNA Extraction procedures start with cell lysis. A buffer that includes Guanidine Thiocyanate or other chaotropes is necessary to mask charges on RNA and water so that the polymer can be purified using solid-phase techniques. Chaotropes linearize the RNA polymer, disrupt hydrogen bonding and destroy the activity of any RNAses present in the cell lysate. Due to their ability to disrupt hydrogen bonding, they also lyse cells by disbanding the phospholipid bilayer.

RNA isolation from cells

To facilitate homogenization, a homogenization column (imaged below) may be used. By passing at high speeds through resins, viscous polymers such as DNA and lipids can be separated and the cell lysate will flow more easily.

RNA purification from protein and carboihydrate

Ethanol is then added to: reduce the overall water concentration in the sample and precipitate proteins. As you would expect, RNA is highly soluble in water. A spin column (solid-phase support, imaged below) is then used to bind the RNA from the cell lysate.

Solid phase RNA extraction

Initially a low chaotrope concentration wash buffer is used to clean the RNA sample and remove proteins while RNA remains attached to the column.

Purification of RNA with ethanol buffers

Next, an ethanol wash removes some of the chaotropic salts that were left over from previous washes.

Finally, without chaotropic salts present, RNAse-free water can be used to elute the RNA sample from the column

RNA Later and RNEasy isolation

RNA Extraction and Purification: Step by Step

Here’s a technique for RNA Isolation and cDNA preparation from Pancreatic Islets. We can use this technique for analysis of gene expression later on. Our strategy is to use solid-phase extraction of nucleic acids from complex cell lysate samples and then we can prepare the cDNA to examine the expression of genes such as insulin.
 
Materials for RNA Extraction and cDNA preparation:
Isolated pancreatic islets
RNeasy minikit (#74104 Qiagen)
QIAShredder columns (#79654 Qiagen)
Omniscript reverse transcription kit (#205110 Qiagen)
Oligo dT primers (dilute to 10 µM; #18418012 ThermoFisher Scientific)
Random hexamer primers (dilute to 20 µM; #N8080127 ThermoFisher Scientific)
RNase inhibitor (dilute to 10 units/µl; #N211 Promega)
 
RNA Extraction/Purification Procedure:

  1. Collect up to 100 islets in an Eppendorf tube and add 350 µl RLT buffer* to disrupt cells.
  2. Vortex thoroughly and add to QIAShredder column with collection tube attached. Spin for 2 min at full speed in microcentrifuge to homogenize the sample.
  3. Add 350 µl 70% ethanol to the lysate and pipette repeatedly to mix.
  4. Transfer to an RNeasy spin column and spin at ≥8000xg for 15 secs to bind the RNA to the column. Discard the flow-through.
  5. Wash the column with 700 µl RW1 buffer and spin at ≥8000xg for 15 secs. Discard the flow-through.
  6. Wash the column with 500 µl RPE buffer* and spin at ≥8000xg for 15 secs. Discard the flow-through.
  7. Repeat column wash with 500 µl RPE buffer and spin at ≥8000xg for 2 min. Discard the flow-through.
  8. To elute the bound RNA, transfer column into a fresh Eppendorf tube and add 30 µl RNase-free water to membrane. Spin at ≥8000xg for 1 min.
  9. To ensure a high RNA concentration, use the eluate to repeat the elution by reapplying to the membrane and spinning at ≥8000xg for 1 min.*
  10. Determine the concentration and quality of the RNA sample.*
  11. To reverse transcribe template RNA into cDNA, prepare the following reaction in one eppendorf:
    Up to 2 µg RNA*
    4 µl of 10X Buffer RT
    2 µl Oligo dT primer
    2 µl random hexamer primer
    1 µl RNase inhibitor
    1 µl Reverse Transcriptase
    Add RNase-free water to take reaction volume to 40 µl.
  12. Mix thoroughly by briefly vortexing and centrifuge briefly to collect reaction to the bottom of the tube
  13. Incubate at 37oC for 60 min.
  14. Place reactions on ice and use up to 1 µl per 10 µl PCR reaction*.

 
Notes for this RNA Isolation Procedure:

  • Step 1. On the day of RNA preparation, add 10 µl of 14.3 M 2-mercaptoethanol per 1ml RLT buffer prior to use. This helps disable RNAses.
  • Step 6. Ensure ethanol is used to dilute RPE buffer concentrate (provided in kit) prior to first use
  • Step 9. Store RNA samples at -80°C if required to prevent degradation
  • Step 10. RNA analysis chip techniques such as Experion (BioRad) use a small amount of sample to provide accurate quantification and assessment of RNA quality
  • Step 11. If RNA has been frozen, thaw on ice to avoid RNA degradation
  • Step 14. Reactions can be stored at -20oC prior to PCR if required

Notes from our readers:

Mr. Young on Google + states:

1. [It is important to] …keep… an amplicon-free, RSase- free, clean environment
2. [Also have] a dedicated space for nucleic acid extraction separate from post amp and Master mix prep

Dr. Carina Jorgensen from Linked In states:

[Check the] integrity of … [your] … RNA … – that is whether or not its degraded – before …[you]…continue with downstream applications…You need to know the quality of RNA to choose your primer for cDNA synthesis, and if you want to continue with real-time qPCR you need to have a certain minimum level when it comes to the quality of your RNA, if you want to be sure that you can trust the following qPCR results. If you want to perform microchip analysis the demand to quality is even higher comparer to qPCR.

Guides for Specific Applications of RNA Extraction on Scigine

HeLA cell RNA Isolation
RNA Extraction from Cumulus Cells
RNA Extraction followed by RT-PCR
RNA Isolation and cDNA Synthesis
Acid-Phenol Extraction of RNA

References

Sambrook, J., and Russell, D. W. (2001) Molecular Cloning: A Laboratory Manual. 3rd Ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
RNA Extraction Methods in Molecular Biology Book Chapter
Bitesize Bio Guide

A Useful Video Related to RNA/DNA Isolation

Immunoprecipitation (IP) Scientific Method Guide

Immunoprecipitation Method Guide on SciGine

Immunoprecipitation Overview

Immunoprecipitation is a method for extracting protein from a solution. Typically, this solution is a cell lysate which you want to analyze. Very frequently you’ll hear your colleagues say, «I’m going to do an IP-pull down on my protein» — when you hear this, you’ll know they’re talking about immunoprecipitation. This technique is a must-have for any biochemist who works with proteins because it’s so versatile. Once you remove a protein from solution, you can analyze it to see what it binds. You can also check out its molecular weight and structure. And, beyond proteins, IP can be applied to RNA and DNA pull downs as well.

In theory, this method is very simple. First, lyse your cells using some sort of lysis buffer. Next, add in an antibody that will bind your protein of interest and form a protein-antibody complex. Then drop in some resin which can bind the antibody. Typically, this resin is either agarose-based or superparamagnetic and is covered with Protein A and/or Protein G. These proteins are specifically designed to bind the heavy chains of antibodies so they can easily pull out your protein-antibody complexes. Agarose beads offer higher capacity per bead but magnetic beads are MUCH easier to separate because you can use a magnet to keep them in place. Finally, spin everything down and remove the supernatant. With the remaining bead-antibody-protein conjugates, you can either denature everything and run a SDS-PAGE western blot, or you can try to analyze your protein’s function with an activity assay, or run it on HPLC or LC-MS, etc. There are so many downstream applications of immunoprecipitation!

Take a look at the attached drawing to understand this method:
Immunoprecipitation Scientific Method
Immunoprecipitation Pulldown Assay
Immunoprecipitation IP Downstream applications

Different Types of Immunoprecipitation (IP) / Pull Down Assays

While the previous description shows the most straight forward and common version of immunoprecipitation, there are many variants of this method. It’s truly versatile and powerful, so let’s take a look:

  • Pre-immobilized Antibody Immunoprecipitation: Instead of adding in antibody for your target and then immobilizing the antibody onto agarose beads, you can pre-immobilize it and then add it into your protein mixture.
  • Co-IP, CoIP, or Co-Immunoprecipitation: In a Co-IP, you pull down multiple proteins along with your protein of interest as a complex. It’s very likely that this protein complex that is physiologically relevant to the signaling cascade that they trigger. As an example, you could pull out clathrin-binding proteins and you’d have a clue as to how these proteins enter/exit cells. Cool right? With this simple technique, you can now map out which proteins interact with each other to cause a certain phenotype!
  • ChIP or Chromatin Immunoprecipitation: In this method, the DNA that is bound to your target protein is what you are after. The idea is to first let your protein bind DNA. Then you crosslink it using formaldehyde and lyse the cells. Next, you fragment the DNA so your protein isn’t bound to ALL the cellular DNA…it’s only bound to the DNA that it interacts with. Then do a regular IP based on this blog post. With this precipitated protein, you can then detach the protein from the DNA using heat and perform PCR to look at what DNA segment is being bound.
  • RNA Immunoprecipitation or RIP: Jut like a ChIP, you can also pull out proteins that bind to RNA inside cells. The strategy is the same as ChIP but you need to use RT-PCR to analyze the RNA that you get. We’ll discuss this technique in the future.
  • Biotin/Streptavidin Immunoprecipitation: Some times, Protein A and/or G, based pull-downs don’t work because your antibody doesn’t bind to them very well (due to protein-to-protein variability). In these cases, you need an alternate strategy. If you can find biotinylated antibodies for your target protein, then you can use streptavidin beads to pull them out! Biotin-Streptavidin interactions are among the strongest molecular interactions known to science.
  • Covalent Capture Immunoprecipitation: If you cannot use Protein A and/or G, and you cannot find the biotinylated antibodies for your target protein, then you have to go back to chemistry. The strategy is relatively simple, but most biologists hate chemistry 🙂 All you need to do is to oxidize your antibody to introduce aldehydes in the backbone. This can be done using periodate oxidation on vicinal diols in the heavy-chains of the antibody. Then using amine-containing resins, you can capture the antibodies via schiff base formation and borohydride reduction. You can also flip this strategy on it’s head and use amine-containing antibodies with aldehyde containing resin.

Some of these strategies are shown here:
Preimmobilized antibody or Coip used in IP Pulldowns
Co immunoprecipitation and Chromatin immunoprecipitation
Steps of Chromatin Immunoprecipitation
Finding DNA binding regions of proteins using ChIP

Immunoprecipitation Scientific Method Step-By-Step

Immunoprecipitation of the Lck protein from T cell lysates has been used to analyse the phosphorylation patterns of this important T cell activation protein.

IP Materials:

Lysis buffer (20 mM Tris pH 7.5, 150 mM NaCl, 1% Nonidet P-40, 1 mM phenylmethane sulfonyl fluoride (PMSF))
Protein G-Sepharose beads (Protein G Sepharose 4 Fast Flow #17-0618-01, GE Healthcare Life Sciences)
Protease inhibitors (HaltTM Protease Inhibitor Cocktail 100X, #78430, ThermoFisher Scientific)
Purified mouse anti-human p56-Lck antibody (#551048, BD Biosciences)

IP Experimental procedure:

Perform all steps on ice to avoid protein degradation.

  1. Pellet cells (107) and resuspend in 800 µl ice-cold lysis buffer containing protease inhibitors. Lyse cells at 4°C for 15 min with mixing.*
  2. Pellet cell debris for 15 min at maximum speed on microcentrifuge at 4°C.
  3. Prepare Protein G-Sepharose beads following manufacturers’ instructions, by washing 1 ml of bead slurry three times in lysis buffer. Resuspend the final pellet in an equal volume of lysis buffer and store at 4°C until use.*
  4. Preclear lysate by adding 50 µl of washed Protein G-Sepharose beads at 4°C with mixing for 1 h. Pellet beads for 20 sec at maximum speed on microcentrifuge and retain lysate in a fresh tube.
  5. Add 1-2 µg of specific antibody (eg. anti-Lck) to lysate for 15 min at room temperature. Add 50 µl of washed Protein G-Sepharose beads and incubate overnight at 4°C with mixing.
  6. Spin for 20 sec at maximum speed on microcentrifuge to pellet immunoprecipitates. Wash complexes three times in 500 µl lysis buffer.*

IP Procedural notes:

  • Step 1. Prepare fresh lysis buffer on day of procedure. If protein is intended to be assayed for phosphorylation levels, also include phosphatase inhibitors.
  • Step 3. Protein G has a high affinity for mouse Ig; for other precipitating antibodies Protein A can be used.
  • Step 6. If samples are to be analysed by SDS-PAGE, add reducing sample buffer directly to washed immunoprecipitates prior to electrophoresis.
  • Protein A and/or G are not great way to immobilize antibodies if doing an IP with serum. Serum contains lots of other Protein A and/or G types which will compete with your IP resin.

SciGine Immunoprecipitation Applications

CDK2 Immunoprecipitation
Phosphoserine Pull-down from G418 clones
Immunoprecipitation and SDS-PAGE of FLAG protein
SRC Kinase Immunoprecipitation and SDS-PAGE
Immunoprecipitation of Rheumatoid Arthritis related RV202

References:

Immunoprecipitation guide by Kaboord et. al
IP Protocol by Carey et. al
ChIP by the Haber Lab

Southern Blot Scientific Method Theory and Guide

Southern blot scientific method title

Southern Blots: Overview & Theory

Southern Blotting is a technique that is used to detect whether you have a specific DNA sequence available in your sample. Most commonly, you will be testing for DNA that you get from cell lysates or after creating your own plasmids. You can also do some amazing things like making transgenic mice and proving that you have selected for certain genes of interest to you! All you have to do is Southern Blot any set of DNA from the mice and you’ll have concrete proof. As you probably understand, this is one of the most common techniques that geneticists and molecular biologists know because everything they do needs to be proved by a southern blot.

To execute a southern blot, first collect some DNA. This can be from a cell lysate or from tissue samples, etc. Next, digest the DNA using DNAse and run these fragments on an agarose gel. This will separate the DNA by size and you’ll know which well in the gel corresponds with which sample. Next, in a similar fashion to Western blots, this DNA is then transferred onto blotting paper (typically made of nitrocellulose or nylon). After the “blotting” process is complete, the DNA is then “probed” using a radioactive (or fluorescent) DNA sequence that is complementary to the sequence that you want to detect. Finally, this “probed” radioactive blot is then imaged using an autoradiograph. The steps for this process are described in the following image:
Southern Blot Scientific Method
Southern Blot Step by Step
Southern Blot Tips and Tricks
Southern Blotting Method

Nuances of Southern Blotting theory

Why is there is a second denaturing step for DNA that’s 15 KB or larger?
It’s because these large pieces of DNA don’t transfer very well to the blotting paper. As you can imagine, the larger a piece of DNA is, the slower it will migrate through an agarose gel. Even after leaving the blotting paper overnight, the transfer may not be complete!

How does the DNA actually go from the gel into the blot?
Through capillary action and wicking! Unlike a western blot where a voltage gradient is utilized to pull proteins into the blotting paper, in Southern Blots, the DNA merely moves over to the blotting paper overnight without much force at all. That’s why it is incredibly important to make sure that the blotting paper and the gel are in close contact. It’s also very important to make sure that the glass plate on top is heavy enough so that it forces the gel and the blot together.

Why is there a UV step?
By using UV after the transfer step, the DNA (which has some free aldehyde groups due to depurination) can react with the nitrocellulose/nylon membrane to form covalent bonds.

What do the NaOH and the HCl do during the denaturation step?
Essentially HCl removes some/all of the purine bases from the DNA and makes the two DNA strands less sticky to each other (because there is less hydrogen bonding). This process is called depurination. NaOH also prevents the two strands from forming hydrogen bonds due to deprotonation of all bases.

How does radiolabeling with P32 work?
Small amounts of DNAse introduce nicks into the single stranded probe DNA. DNA polymerase then utilizes the dATP32 from solution to repair these nicks and incorporates these radioactive phosphates into the backbone of the DNA.

Southern blot Step-By-Step Guide

Materials for Southern Blot of Mouse Tail DNA

Denhardt’s Solution 50X (5g Ficoll 70000, 5g Polyvinyl pyrrolidone, 5g BSA Fraction V in 500 ml water)
Hybridization cocktail (25 ml 50% dextran sulfate, 25 ml 20X SSC, 50 ml formamide, 1 ml Tris 1M, 2 ml Denhardt’s Solution 50X, 1ml 10% SDS) – prefilter before use
TE buffer (10 mM Tris HCl pH 7.5, 1 mM EDTA)
BamHI enzyme and buffer (#R0136S New England BioLabs)
100X BSA (B9000S New England BioLabs)
Gel loading dye (#G2526 Sigma)
TBE buffer 10X (#93290 Sigma)
SSC buffer 20X (#S6639 Sigma)
[α32P]dCTP (3000 Ci/mmol; #NHG013H250UC Perkin Elmer)
Sephadex G-50 (#G50150 Sigma)
Ready-To-Go DNA Labeling Beads (-dCTP) (#27-9240-01 GE Lifesciences)
Whatman paper (#WHA10427810 Sigma)
Salmon Sperm DNA (#15632011 ThermoFisher)

Southern Blot Protocol
DNA digest and gel electrophoresis

    1. Setup DNA digest reactions from collected tail samples as follows:

10 µl DNA (12 ug)
6 µl 10X BamHI buffer
0.6 µl 100X BSA
1 µl BamHI
42.4 µl H2O
Incubate at 37°C for at least 5 hours.

  1. Add 6 µl gel loading dye to each sample.
  2. Run samples of a 0.8% agarose gel in TBE for 16 h at 25V.
  3. Stain gel in 1 µg/ml ethidium bromide for 30 min.
  4. Take a photo of the gel.*

Transfer

    1. Soak gel in 0.2 N HCl for 10 min with shaking to depurinate (remove purines from DNA). Rinse with water.
    2. Soak gel in two washes of 500 ml 0.4 M NaOH/1.5 M NaCl solution for 30 min each time.
    3. Soak gel in two washes of 500 ml 1 M Tris/1.5 M NaCl solution for 30 min each time.
    4. Setup transfer stack by placing 2 layers of Whatman paper on glass plate (wet before laying down) with soaked gel on top (flipped over). Cover with plastic wrap.
    5. Using a razor blade, cut out plastic over gel and place wetted membrane on gel. Place two layers of Whatman paper on top (wet before laying down).
    6. Place a stack of paper towels on the top and cover with a glass plate and transfer in 6X SSC overnight.
    7. Dismantle and air-dry membrane.
    8. Expose membrane to UV for 90 s (DNA side down). Bake between Whatman paper at 80°C for at least 1 h.

Labelling the probe

      1. Denature probe by adding 2 µl probe (25-50 ng) to 44 µl TE and incubating at 95°C for 4 min). Immediately place on ice.
      2. Add 46 µl denatured probe and 4 µl of [α32P]dCTP to Ready-to-Go labelling bead tube. Flick to mix.
      3. Incubate at 37°C for 15-30 min.
      4. Add 50 µl TE and run through a G50 sephadex column

Probe Hybridization and Autoradiograph

      1. Add 50 ml hybridization cocktail to the membrane.
      2. Add 300 µl of 10 mg/ml salmon sperm DNA to labelled probe and denature at 95°C for 5 min. Immediately place on ice. Add to membrane and cocktail solution.
      3. Hybridize overnight at 42°C.
      4. Wash briefly with 50 ml wash solution (0.2X SSC/0.1% SDS).
      5. Wash for 30 min with 100 ml wash solution at 50°C. Repeat.
      6. Check blot with Geiger counter to estimate cpm.
      7. Expose membrane to film and store at -70°C.
      8. Develop after 2-5 days.

Notes on this Southern Blot method

      • During DNA digest Step 5 you can note the position of the marker bands by using a fluorescent ruler or marking bands directly on the gel by punching small holes with a needle.
      • This method uses dCTP32 and not dATP32 as we talked about in the theory section
      • After undergoing HCl and NaOH treatment for the denaturation step, it is very important to neutralize the gel once again.
      • If you don’t want to buy the labelling beads, an alternative strategy is presented here at MIT Cores

Application of Southern Blots on SciGine, a Scientific Method Search Engine

References:

ASU Southern Blots Guide
MIT Core Guide
DNA Blot from Davidson College

Southern Blot Video by Shomu’s Biology

HPLC: Biochemical Analysis. A Step-By-Step Method Guide

HPLC Analysis Step by Step

HPLC Method Overview

HPLC, or high performance liquid chromatography is an amazing analytical technique for chemical compounds including biopolymers, small molecules, and polymers. In this method, a sample is first dissolved to make a solution. This solution is then injected into a “column” that contains resin that will interact with the sample. This will slow down the movement of the sample through the “column” and as the sample comes out the other side of the column, it is detected. This allows you to know both the time at which the sample comes out and the intensity of the sample that was detected. Here’s an overview of this technique:

HPLC Bioanalytical Method Guide

So, while there is continuous flow of some buffer through the column, we also inject our sample and observe as different molecules within the sample come out at different “retention times”. The detector on the end of the column can be any kind of detector but the most common types are refractive index (RI), ultraviolet (DAD), and fluorescence (FLD). Each of these will detect different properties of the molecules that come out of the column and display a chromatogram.

HPLC Chromatogram Guide

Types of Chromatography

Different column resin compositions determine the kind of chromatography that you are running and what molecules you can separate.

  • Normal Phase: The column is filled with silica particles which are polar and the buffer running through the system is non-polar. Once you inject your sample, polar particles will stick to the silica more and have a longer retention time than non-polar molecules.
  • Reverse Phase: The column is filled with hydrophobic particles (actually they are silica particles with long hydrocarbons on the surface). The buffer that is running through the system is polar (such as acetonitrile/water or methanol/water mixtures). This means that hydrophobic molecules will stick to the resin more and be retained longer.

Complete Step by Step HPLC guide

Materials

HPLC autosampler vials I only use autosamplers since manual injection is tedious 🙂
Centrifugal filters with 0.2 um pores To clean up samples
Eppendorf vials For centrifuging
HPLC machine

Methods

In a typical HPLC procedure you can decide the following variables:

Variable What it does
Flow rate With fast flow peaks come out sooner but there’s they’re harder to resolve and tend to blend together. For more resolution, run slower.
Pressure Affected by flow rate and solvent
Solvent Buffers Determines signal intensity, how quickly the peaks come out, signal fidelity
Column Type Determines the type of interaction with the sample
Detection Parameters If using UV or FLD, you need to set the right excitation/emission wavelengths

Since HPLC is a very machine-variable technique, I can only provide general guidelines.

For sample preparation:

  1. Dissolve your biopolymers or small molecules in a suitable solvent such as methanol
  2. Centrifuge at 10,000 rcf in an eppendorf vial and keep the supernatant to remove any large particular matter
  3. With a centrifugal filter, add 500 ul of your sample solution onto the top
  4. Centrifuge at 10,000 rcf and collect the filtrate (the solution that successfully passes through the filter)
  5. Load this sample into an HPLC vial

For setting up the HPLC machine:

  1. Make sure you have all your buffers set up
  2. Open the purge valve and purge the system for 5 minutes.
  3. Add your samples into the autosampler tray
  4. Stop the purge
  5. Close the purge valve
  6. Run the system at a normal flow rate (1 ml/min) with your buffer to equilibrate the column for 10 minutes
  7. Make sure that your pressure is stable (ie, less than 2-3 bar of fluctuation)
  8. Set up your sequence and your method
  9. Run a standard before your actual samples or as part of the same sequence

Example buffer system to determine Fluoresceinamine levels in samples:
Sample: Add 10 ug of fluoresceinamine into 1 ml of Acetonitrile.
Buffer: Pure acetonitrile buffer on a C-18 column; this is “reverse phase”.
Flow rate: 1 ml/min.
Column: 4.6 mm x 30 cm size.
Detection: Detect via a fluorescence detector set to Excitation @ 485 nm and Emission @ 535 nm.

Notes on HPLC methodology

  • To clean the system and equilibriate it, you need to run enough solvent. However, this amount varies column-to- column. A typical 4.6 mm x 30 cm column should be clean when you follow the procedure above.
  • Isocratic means that the solvent concentration stays constant throughout the run.
  • It is useful to run standards before your samples as well as with your samples. Standards make it easy to identify which peak pertains to your molecule of interest.
  • Always use HPLC grade solvents. This is especially true for solvents like THF which are frequently sold with inhibitors that also complicate your ability to detect your molecule of interest.

Applications of HPLC on SciGine

HPLC is such a versatile technique. Take a look at these methods on SciGine which assay different types of chemicals in various samples.

References

ChemGuide Summary of Technique
Method Guide from Waters
Overview on Wiki

Using PCR To Amplify DNA, A Step-By-Step Guide

PCR Method Guide on SciGine

PCR Biological Method Overview

PCR, or polymerase chain reaction, is a method to amplify a segment of DNA for analysis. Because it is such a powerful technique, there are a HUGE number of situations where PCR may be used. Some common reasons for using it are:

  1. Microbiology: You need to know if your bacteria was transformed properly with your plasmid
  2. Oncology: You need to know if a particular gene exists in your cancerous cells
  3. Forensics: You need to know if a certain criminal’s DNA was present at the crime scene

The basic steps of PCR include:

  1. Designing primers to designate a target DNA sequence to amplify
  2. Mixing together the primers with the target DNA strand, polymerase enzyme, and deoxynucleotides
  3. Running a thermocycler multiple times in “cycles” to repeatedly:
    • Separate DNA strands
    • Allow primers to anneal
    • Allow the polymerase to attach and synthesize a new DNA chain
  4. Sequencing or Electrophoresis to prove that the PCR worked

In a nutshell, here is what Step 3, above, looks like:

PCR, a biological method, amplifies DNA - SciGine

Ingredients in a PCR mixture – What every component does

In general, a PCR tube will contain the following items:

Water: Solvent
dNTPs (or deoxynucleotide triphosphates): Single bases A, T, C, and G which are used by the polymerase while replicating the DNA. As the polymerase adds base pairs onto the new DNA strand, one base pair is used at a time.
MgCl2: An essential cofactor for the polymerase enzyme
Primers: Short segments of single-stranded DNA used to frame the DNA region that needs to be amplified. They are complementary to the template DNA strand only at defined locations around the target sequence.
Target DNA: The DNA “template” that you want to make copies of. This can be a full DNA chain or a part of a longer chain.
Taq Polymerase: An enzyme from Thermis aquaticus that uses dNTPs and replicates DNA starting from the 3′ end of a template strand towards the 5′ end. Taq  is used in PCR specifically because it is resistant to the high temperatures used for separating DNA strands during Step 3a above .

Complete Step-By-Step PCR Protocol

Materials

PCR tubes

PCR temperature Cycler

Pipettes

Reaction Buffer

10 mM Tris-HCl, 50 mM KCl, and 1.5 mM MgCl2, pH 8.3

Methods

  1. For each PCR tube set up the following mixture of materials up to a 50 ul total volume on ice

    dNTP solution

    200 uM for each dNTP

    Forward Primer

    0.2 uM

    Reverse Primer

    0.2 uM

    Target DNA

    Less than 1000 ng

    Taq Polymerase

    1.25 units per tube

    Nuclease Free Water

    Q.s up to 50 ul

    Mineral Oil

    Add a little bit on top of each tube if you don’t have a heated lid on the temperature cycler

  2. Pipette each tube up and down several times, gently, so as not to add bubbles
  3. Centrifuge each tube down for 1-2 seconds at 100 rcf to bring all contents to the bottom
  4. Heat up the temperature cycler to 95oC
  5. Quickly transfer tubes from ice to the temperature cycler and begin thermocycling
  6. Typical thermocycle procedure:

    Step Name

    Temperature

    Time

    Denaturation

    95 oC

    30 seconds

    Cycle Denaturation

    95 oC

    30 seconds

    Cycle Priming

    50-60 oC

    60 seconds

    Cycle Extension

    72 oC

    1 minute per kilobase of target DNA

    Final Extension

    72 oC

    5 minutes

    Hold Temperature

    4 oC

    Infinite

  7. Repeat the above “cycle” steps 35-45 times.
  8. Make sure to keep the samples on ice while you are not using them. The infinite hold sequence allows you the flexibility of leaving your samples in the temperature cycler while you are busy with other lab work.

Notes on this PCR Methodology

  • Note 1. It is important to design your PCR primers to be specific to only the regions flanking the target sequence. Typically, specific primers are ~30-40 bases in length.
  • Note 2. The Tm (melting temperature) of the primers affect the temperature in Step 3b and the “Cycle priming” step.
  • Note 3. For greater accuracy/fidelity while copying DNA, a Pfu polymerase (from Pyrococcus Furiosus) may also be used.
  • Note 4. With 35 cycles, the target DNA is amplified 236 times its initial concentration. For times when you have very little target DNA, you can amplify 45 times or more.
  • Note 5. For Primer design, you can use tools such as Primer 3. Tools are also available for calculating the Tm of your primers such as this Tm calculator.
  • Note 6. G-C rich sequences will need longer denaturation times (typically up to 5 minutes) because of the additional hydrogen bonding. Any template with more than 60% G-C bases is considered G-C rich.