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1)      His SpinTrap (GE Healthcare, Singapore)

2)      Bio-Rad Protein Assay (Biorad, Singapore)

3)      InstantBlue Protein Stain (Expedeon, Singapore)

4)      Trans-Blot Turbo Transfer System (Biorad, California, USA)

5)      Anti-6X His tag antibody (Abcam, Cambridgeshire, England)

6)      Goat anti-Mouse IgG (H+L) Secondary Antibody, HRP (ThemoFisher Scientific, Waltham, USA)

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I think I will stop using this interface because no one really has anything for me here. The post below is my last post for this module.

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I think I will stop using this interface because no one really has anything for me here. The post below is my last post for this module.

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*update* I'll stay on till the end of the Molecular part.

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The last practical (for molecular cloning), we carried out a cycle sequencing with 4 independent primers that flank externally and internally of the insert. I would want you to consider why do we carry out the reaction this way? As in why can't we have 2 primers in the same reaction?

Cycle sequencing is similar but NOT IDENTICAL to PCR. PCR expands amplicons exponentially while cycle sequencing, linearly. Cycle sequencing also uses another special Taq polymerase that has two further mutations - to remove the 5' - 3' exonuclease function & to pick up ddNTPs as similar probability as the dNTPs. ddNTPs has a proton replacing the 3'-OH groups, which halts elongation during cycle sequencing phases. The ddNTPs itself contains a fluorescence and will light up during sequencing activities (with an appropriate wavelength). I.e., each ddNTPs (G/A/T/C) contains a unique fluorescence colour that will be captured by the sequencer and interpreted by the program attached to the sequencer, usually an independent computer with an appropriate program installed.

Sometimes we can experience issues such as compression in sequencing. This happens in high GC, repeating content. And it forms 3' hairpins that may cause these fragments to run similar rates as normal, smaller linear fragments during sequencing phase. As such, the sequencer may not be able to differentiate between nucleotides, and may interpret it as an error. To tackle this, we either use (1) high temperatures and formamide during electrophoresis, (2) bisulfite treatment, or (3) replacing dGTPs with dITPs or 7-deaza-dGTP. Bisulfite treatment requires you to conduct sequencing with the original and treated strands to detect truncation in the former. That way, we can redesign primers or parameters to conduct sequencing better. dITPs and 7-deaza-dGTP has one side group lesser. I.e., they cannot hold 3 hydrogen bonds (with dCTP) as compared to the original dGTP. And in this manner, we can use lesser heat energy to linearize the strands.

After the sequencing, we would need to analyze the data and check whether it will be feasible to transform it into another E. coli strain called BL21 (DE3). This is an expressional cell line that will transcribe your insert's content. We will then categorize the protein produced in practical 5 - 8.

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We all know that we are using a directional cloning. And this would mean that we use two independent enzymes to cut and force the orientation of the insert of our liking.

Now consider that you are using a T-A or blunt end cloning. How do you determine the orientation of your insert now?

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Prior to this practical, we have to count colonies and pick 4 random ones for analysis. Bear in mind that colonies growing on the LB+Amp plate represents a bacteria colony which contains the pET11a plasmid. But it DOES NOT tell us whether the plasmid contains the insert or not. Merely because the backbone itself has the beta-lactamase gene (not the insert).

We isolate the plasmid with the miniprep kit. Because it is a low copy plasmid, we need to increase the pellet size to get more copies of the plasmid. We resuspend in resuspension buffer. This has Tris-HCl (as a simple buffer ~7.4), RNAse and EDTA. We use RNAse because we don't need them (we want the plasmids only right?). It also reduces the viscosity of the cytosol; nucleotide sequences (if in abundant) can cause viscosity of lysate. EDTA is a metal chelator that reduces the chance of DNAse activation.

Lysis and neutralization steps are very crucial. If we prolong the incubation, there may be higher chance of nicks to form on the plasmids. As such, vortexing is prohibited to prevent shearing of DNA material. Lysis buffer contains SDS and NaOH. SDS is to lysis the hydrophobic membrane and denature the proteins. NaOH denatures the DNA material into singular strands. Alkaline protease helps to deactivate most proteins. Neutralization solution contains sodium acetate. NaAcet reverses the pH so that the DNA material will find its own complementary sequences again. Smaller fragments like plasmids will be able to come back easily as compared to genomic DNA, which takes a longer time to complement properly. Usually they will mis-complement and form aggregates/precipitates (together with the proteins or debris) which you saw in the practical.

After the spin down, you transfer supernatant (together with the plasmids) into the column. The column works in similar fashion as the cleanup columns. The difference is that we do not need to use chaotropic agents to help DNA bind to the silica matrix. High salts are necessary to keep the plasmids linked to the matrix. Washing solution contains ethanol, which helps to wash the DNA material. Finally, we elute out with water (with low/no salts).

We digest with NdeI and EcoRI again to test for presence of insert. The keyword is "presence". We fit in a 1kb insert into the vector and re-digest it with the same set of enzymes for testing. We assume that what goes in should come out in the same way. But if we think carefully, a 1kb band can be represented by random sequences as well. So we can only assume that the 1kb may be our gene of interest. We would need at least one more test to confirm this statement, and that is in practical 4 - sequencing.

Plasmids have a strange way of presenting itself on the agarose gel (with DNA stain). They usually feature three bands - a supercoil (fastest), linear and relaxed (slowest). If your technique is good enough, you usually see the supercoiled only. Plasmids also have a strange way of running on the gel. If you increase the amount of DNA stain (in the gel), the plasmids will run slower at first, and then faster again. This is correspondent to the orientation of the twist of the plasmid. Just imagine a rubber band (looking from the top of it) and twist it clockwise. Continue to do this until the rubber band is shrink into a small ball. This is a supercoiled (let's say negative-oriented supercoil). When the DNA stain is intercalated in between the grooves, it unwinds the DNA by rotating it anti-clockwise. As you increase more and more stains, the anti-clockwise twist increases until you return back to a circular form. Continue to add stains and the twist will now turn the other way back into a supercoiled again (a positive-oriented supercoil).

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Alright guys (and gals), you have already seen each others' colony numbers. What you can do is to have vector:insert ratios 1:1 as one group, while 1:2 as the other. Use a statistical program that you are familiar with to check for significant difference.

The simpliest one I can think of is this webapp - https://www.graphpad.com/quickcalcs/ttest1.cfm

Key in the details for a student t-, two-tailed test. Do not place in group 46's results because it is not valid; they did not get any colonies.

Then ask yourself - if it is significantly different, what does it mean; if it is not sign. diff. then what can we say? We can discuss this tomorrow.

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Dear all,

Here are the colony counts for all of the groups:
vector:insert
Group                                1:1                              1:2
46                                         0                                  0
47                                      581                             928
48                                       68                               189
49 887 258

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Hi Johann,
Referring to page 1-9 of the protocol, step 3, 'Incubate the tubes at 72 degrees for 2 minutes', I remembered you mentioned that this step was used to denature the primers. I am confused as to why we have to denature the primers right after adding them in step 1 and before step 6 (when Reverse Transcriptase was added)?

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Today, we took  the digested materials - RT-PCR & pET11a vector - and underwent a cleaning process. The DNA materials were digested with sticky ends so as to control the direction of the sequence. I.e., we apply a directional cloning technique. This is very essential for experiments that require expression of proteins. There are two other types of cloning processes but we did not apply them today; the other two types are T-A and blunt cloning. Blunt end cloning have a very high chance of self ligating, but can be minimized with the help of a phosphatase. The most common ones on the market are the calf intestinal phosphatase (CIP) or the shrimp alkaline phosphatase (rSAP).

Some researchers do not wish to use phosphatase because the latter binds very strongly to DNA molecules. And at times, downstream processes can be affected by the presence of the phosphatase. The other reason is that there is a chance for mutations to occur. When phosphatase is applied to the vectors, their 5' ends are unable to undergo ligation. But the insert can still be ligated at the 3' ends. I.e., there will be two nicks where the 5' phosphate groups were removed. After transformation, the cell will use its repair system to replace the nicked nucleotide. However, there is also a chance of it being mutated due to the error rate. As such, mutations can still occur.

The pieces were then "glued" together by a T4 DNA ligase, which makes use of ATP to actively combine the 3'-OH group to the 5'-P group. Do bear in mind that the 5' phosphate group is the second phosphate. Ligase can be affected by temperature and length of time. The most common condition researchers usually use is a 12-16 degrees Celsius, overnight.

We applied a chemical transformation which makes use of Ca²⁺ ions and a large thermocline to induce transfer. Calcium ions would probably stay near the cell surface where some negative proteins reside, to destablize and increase permeability. Similarly, it also draws the plasmids closer to them because of the negative DNA. During heat shock, the jump from 4 degrees Celsius to 42 degress Celsius provides a large enough impact to allow DNA material to be "sucked" in via the open pores. Another method of transforming is to use an electrical method - electroporation. The method requires a current to accelerate the DNA particles to forcefully puncture the walls and enter the cell; the cell will fix its walls during recovery stage. But this method must not have any salts available, at least to the minimum. Because salts can cause an arc/spark to form, and this will kill the cells instantly. Therefore, we need to dialyse the ligation reaction prior to electroporation.

Recovery steps are very important, not because the cells need time to recover from the heat, but to produce enough beta-lactamase to survive on the LB+Amp plate. The Ap gene that is in the manual produces the bla proteins, which corresponds to the beta-lactamase. The beta-lactamase is necessary to cleave the beta-lactam ring of the ampicillin, rending the latter useless. I.e., the colony will virtually have its surrounding cleared off of ampicillin. We must also be very careful in incubating for too long because it may allow satellite colonies to grow around the initial one. Satellite colonies DO NOT have the plasmids at all.

Now, what does it mean if there are colonies grown on the LB+Amp plate?

Just additional questions for you to ponder - How will the results look like if we applied T-A cloning instead? What is an essential point to take note if you want to express proteins in T-A cloning? How does the colonies form as it is in T-A cloning?

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Let's discuss what we have covered in Practical 1. We all understand that the total RNA was extracted from NIH 3T3 mouse fibroblast, reverse transcribed and then amplified. The reverse transcription reactions were split between transcriptase and one without transcriptase. The reason why we do this is to use the latter as a negative control. The gene that we are amplifying is the Lactate Dehydrogenase A gene (LDHA). We use degenerate primers (with special sequences) to anneal and flank the gene of interest (GOI). The primers have the restriction enzyme sites - NdeI & EcorI - and 6x histidine sequence embedded. We also added extra base pairs at the end of the primers to make sure that the REs can sit comfortably onto the sites. Because in vitro settings, there are no external factors to help keep the dsDNA intact. So the entropy at the ends of the dsDNA tend to be very high. I.e., they tend to "open up" despite being complementary. So, we usually add extra base pairs just in case.

After the RT-PCR, we have to test whether the tubes have any bands or not. I.e., whether tubes 1 and 3 have bands at the right sizes. So technically, we need to do a gel electrophoresis prior to the clean up. But to expedite our practical, we let you students carry on with the clean  up together with the run. So just bear in mind that this is not the usual practice; we have already optimized it to allow you to carry out in this way.

Alright, remember that the clean up consists of chaotropic agents that will bind to the dsDNA. But it's not at all locations - just know that there are some parts of the DNA that it will bind to. So this creates an "arm" for it to hold onto the silica membrane when the former is applied to the latter. I.e., the chaotropic agents will bind to the silica, in which, the agents are holding onto the DNA. The washing buffer, consisting of ethanol, will wash the pellet and eventually eluted out with deH₂O. Why don't we use elution buffer (with EDTA) is because EDTA is a metal chelator. What it does is to "hug" onto metal ions and don't let reactions use them. And we know that enzymes like polymerase will need to use metal ions as co-factors. Therefore, the presence of EDTA can suppress/prevent reactions to take place, and researchers prefer to use water instead.

At the end of the practical, we use enzymes to digest at certain places on the DNA strands. What I want you to do is to go back and study how the cuts are being performed. And also consider why did we use BamHI for the vector? (I know this has been featured in the lecture notes before, so you will definitely know the answer).

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DNAseI and polymerase - Why MgCl2 and MnCl2 have various outcomes?

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DNAseI and polymerase - Why MgCl2 and MnCl2 have various outcomes?

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Just so you all understand - what I am going to discuss here is entirely additional information. So you don't really need to read or memorize this in-depth.

Mg2+ and Mn2+ ions act as cofactors for the said enzymes. Mn2+ is probably interacting with the DNA and enzyme in a way that destabilizes the environment between both of them. But this may be with the help of the size of the metal ions as well, causing steric hindrance.

For DNAseI with Mg2+, only one of the two double strand DNA is close to the enzyme's active site. Whereas for Mn2+, it allows strands (at different periods of time) to be cut at similar location, presumably at the N7 of a guanine and an adjacent phosphate.

If you are interested, you can read this article - http://www.jbc.org/content/255/8/3726.full.pdf

Now, for polymerases with Mg2+, it provides specificity in the nucleotide incorporation during elongation between 1 - 4 mM. Whereas for Mn2+, it increases the amount of non-specific incorporation during elongation, which entails to increasing chance of mutations.

So why does this happen? Polyermases have an affinity for nucleotides. But not all nucleotides have identical affinity, which forms different Km values per nucleotide. Mn2+ has a strange phenomenon to alter the Km of the nucleotides to the enzyme. As such, less specific nucleotides may be incorporated.

If you are interested, you can read this article - http://www.pnas.org/content/81/23/7378.full.pdf

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Hi all,

So some of you may not be as clear of why there are forward and reverse primers. Here is the animation that I recommend watching. If you are still unsure, let's discuss again next time.

https://www.youtube.com/watch?v=eEcy9k_KsDI

Someone also asked on how fast a typical polymerase expands? Usually it expands roughly 1kb per min, but recent polymerases are more advance, and can expand much more with the same stipulated time. I would say that the rule of thumb is 1kb per min. Bare in mind ALSO that PCR is an in vitro process, and lacks other factors. I.e., in vivo expansion may jolly well be factors times faster than 1kb per min.

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TRIzol reagent (Invitrogen, Singapore) – Practical 1

First Strand cDNA sSynthesis Kit (ThermoFisher Scientific, Singapore) – Practical 1

FavorPrep PCR Clean-up Kit (Favorgen, Taiwan) – Practical 1 & 2

DreamTaq DNA Polymerase (ThermoFisher Scientific, Singapore) – Practical 1

GeneRuler 1kb DNA ladder - (ThermoFisher Scientific, Singapore) - Practical 1 & 3

T4 DNA Ligase (Promega, Madison, USA) – Practical 2

NdeI (NEB, Ipswich, USA) – Practical 1 & 3

BamHI (NEB, Ipswich, USA) – Practical 1 & 3

EcoRI (NEB, Ipswich, USA) – Practical 1 & 3

Wizard Plus SV minipreps DNA Purification Systems (Promega, Madison, USA) – Practical 3

BigDye Terminator v3.1 Cycle Sequencing Kit (ThermoFisher Scientific, Singapore) – Practical 4

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The main purpose of this interface is to have a platform where we can all discuss about the practical's concepts. If you have any doubts/questions, please post it up here ya? Please do not post any inappropriate things, or flame anyone in this discussion.

You can also use the "attach file" function to take photos of your drawings if necessary.

You can also use your mobile as an alternative.