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Our lab uses old Gilson Pipetman pipettes. I've been using some for making mRNA by in vitro transcription and started running into RNAse issues. I'm trying to clean everything up, including the pipettes. I'm not sure these old pipettes can be autoclaved, at least not in one piece, I think I can take them apart and autoclave the parts that might touch the solutions. Is there some way to wash the pipettes out effectively and remove any RNAses?
I use boxes of RNAse free barrier tips, do all my work on new pieces of aluminum foil, use commercial RNAse free water, gloves, labcoat, facemask, etc. However yields from the in vitro transcription kit have been reduced, and yields from the tailing kit are even worse, sometimes losing 90% of the RNA. There's only so much I can do, our building is old and full of dust and the damn pipes above my work bench leak ( all the pipes leak, had 2 major pipe bursts in the last 6 months ).
I'll probably have to get new kits. Just have to prove to my advisor that the kits are contaminated.
According to the Gilson Pipettman User Guide you can autoclave the following parts: Tip ejector, Tip holder and the connecting nut (have a look into the PDF linked above if you are not sure about the parts). All other parts can not be autoclaved. Gilson says the following conditions can be used (page 17 in the user guide): Autoclave for 20 minutes at 121°C, 0.1 MPa.
However, when you really have a DNAse contamination, you will not get rid of it with autoclaving, since RNAses are too stable. You can either try the various solutions on the market which made for killing RNAses or simply get a new set of pipettes used only for RNA work (on a different bench in a different lab with a completely different and fresh set of chemicals and plastic ware). And I would recommend filter tips for this work. They are more expensive but in the long run it will save money since you don't have to repeat experiments.
Regarding your leaking pipettes: The pipettes contain rubber rings and teflon seals in the inside which have to be replaced from time to time. The image below shows how (parts C and D), the ring can be purchased from Gilson. If you don't know how to do this work (and have no technician who does) you can also send in the pipettes for service. This costs a bit of money, but is ways better than to use unreliable, leaky pipettes.
And don't worry, Gilson pipettes are pretty good. I have used now more or less all brands on the market and I still prefer them. They only need some service from time to time.
Prepare the Sample Tubes
You will use 1.5 mL tubes to extract the DNA samples from saliva.
To start, prepare each tube by labelling them with a permanent marker.
Even if you only have one sample, it’s good practice to label the tube clearly. For example, if the sample is from a person, you could use their initials. It’s also a good idea to mark the date of the sample.
Prepare Saline Solution
You will need salt water (saline solution) as a mouthwash to collect your cheek cells.
In a small glass or similar, mix a pinch of table salt with water. A shot glass is perfect for this. The measurements do not need to be exact. A pinch of salt for a small sip of water is a good rule of thumb.
You do not need to make much – a small sip is enough.
Why the salt water? In this protocol, the aim is to get a sample of DNA from cheek cells. Your saliva, after rinsing your mouth will naturally contain cheek cells, which will be broken open during the protocol to release the DNA. The salt, i.e. sodium chloride, is used to stabilise the DNA, once it has been released.
Rinsing your mouth
Pour the salt water into your mouth. Don’t use too much, just a small sip is enough. Rinse your inner cheeks vigorously for 30-60 seconds. When you are done, spit the saline solution into a new glass. (or, if the shot glass you used to mix the salt water is empty, you can reuse it).
The goal of this step is to loosen as many cells from your mouth as possible. You can use your teeth to gently scrape your cheeks and tongue while you are swirling the salt water around in your mouth. You can also touch your inner cheeks with your tongue. Careful to not hurt yourself – there’s no need for blood, just saliva with lots of cheek cells.
Transfer your sample into the microcentrifuge tube
For this step, you will use your saliva sample (1), the microcentrifuge tube you labelled in the beginning (2), and a transfer pipette (3).
Use the transfer pipette to transfer your saliva sample into the microcentrifuge tube. Fill it up to the 1.5 mL mark.
It is time to use the centrifuge. This will use gravitational force to concentrate the sample.
Put the centrifuge tube with your saliva sample into the centrifuge. Make sure to balance the centrifuge with another sample or with another counter weight.
If you only have one sample, the easiest way to balance the centrifuge is to fill another tube with water and use it as a balancing tube.
Using the centrifuge in an unbalanced way is dangerous and will break the device. Follow our tips for balancing a centrifuge in the manual here. In this case, for example, you could either use a second sample as a counter balance or fill up another tube with water. Tubes must always be balanced with another tube of equal weight.
Once the sample tube is balanced in the centrifuge rotor, close the lid and activate the centrifuge module. Set the speed to 4,000G and spin for 90 seconds.
Recovering the Pellet
Check the sample tube after centrifugation has finished. All the cheek cells should now be concentrated in a small white ball at the bottom of the tube (1). This is called a pellet. The remaining liquid (2), called the supernatant, should be clear.
In this step, you will remove the supernatant, so only the white pellet remains.
Check that your pellet is firmly attached to the bottom of the tube. If it is, you can carefully pour the supernatant away.
If the pellet is not firmly attached to the bottom of the tube, try spinning the sample again in the centrifuge to attach it to the bottom of the tube. If it remains loose, you can use the micropipette with a fresh tip to slowly transfer the supernatant out of the tube. You can also try using the transfer pipette you used earlier, but it might be difficult to control and could end up disturbing the pellet.
Resuspending the Pellet
You should now have a white pellet in your sample tube. It should be about the size of a matchstick head.
If your pellet is smaller than a matchstick head, you may not enough cheek cells to get a concentrated DNA sample. In that case, go back to step 2 to concentrate additional cheek cells from saliva. You can use the same sample tube and simply add more sample to the existing pellet.
Once you have a large enough pellet, you can resuspend the cells into the remaining liquid that is still in the tube. You now have a concentrated cell sample in a small volume of liquid.
Make sure the tube is closed, then mix the cells from the pellet into the liquid by flicking the tube. The cells of the pellet are now resuspended in the liquid.
In this next step, you will use the micropipette (1) to transfer the resuspended sample (2) into a 0.2mL PCR tube (3), so that you can heat it in the thermocycler.
First, set the adjustable pipette to the maximum volume of 20μl.
Make sure the pipette has a new pipette tip. Then use the pipette to transfer the cell mixture of the sample to the 0.2mL PCR tube. Carry on until the PCR tube is almost full, or until you have no sample left. Add as much of the cell mixture as possible to the PCR tube.
Labelling the PCR Tube
Finally, click the lid of the PCR tube closed and label the tube to identify the sample, similarly to the centrifuge tube.
Label the side of PCR tubes, not the lid. The PCR machine has a heated lid, so any ink on the tube lid might come off.
Heating the sample
In this step, you will use the thermocycler as a heat block to boil the cells and burst them open, to release the DNA into the solution.
Place your PCR tube with your sample cell solution in the thermocycler block.
Set up the thermocycler to heat the sample at 99°C for 10 minutes.
Mixing the Sample
After heating the sample for 10 min, we will prepare it again for centrifugation.
First, take the PCR tube out of the thermocycler block.
The block and the heated lid will still be hot, so take extra care.
Flick the PCR tube for 5 seconds to mix the sample.
Centrifuging the sample
In this step, you will spin the sample to separate the supernatant from the cell debris. Now the cells have burst thanks to the heating step, the DNA will be released from the cells and floating in the supernatant.
The molecular weight of DNA is lighter than the other cell material, like proteins and cell walls. By spinning the sample with centrifuge, we seperate the cell material from the DNA, which gives us a cleaner DNA sample.
To spin the PCR tube with your sample (3) in the Bento Lab’s microcentrifuge, you will need to use the PCR tube adapter (1) that sits in a normal microcentrifuge tube (2) and converts it to fit a PCR tube.
Remember to balance your centrifuge. So, if you are only working with one sample, prepare another PCR tube with an amount of water equivalent to your sample.
Set the centrifuge to run for 90 seconds at 8kG.
If you need help operating the Bento Lab centrifuge, check the user manual. Once you the lid is closed, select the time mode (1). Set the force (2) and time (3) before confirming.
Cleaning up the sample for storage
After centrifugation, all the cell debris has been forced to the bottom of the PCR tube (1), leaving only the DNA in the liquid supernatant (2). The supernatant should look clear, like water.
Finally, you will transfer the supernatant into a new PCR using the micropipette.
Set the micropipette to 20μL and put on a new tip. Transfer 40μL of the clear supernatant into the new PCR tube.
Be careful to avoid pipetting any cell debris into the new tube. You should only transfer the clear liquid supernatant. Avoiding any of the cell debris will reduce the chance of interference with the DNA sample.
Labeling and storage
The new tube now contains only the DNA in the liquid. It is called the template sample, and can now be further used for analysis using protocols like PCR.
Label the tube again, so that you can identify which template sample it is.
Finally, if you are not using the template sample in another protocol right away, store it in the freezer at around -20°C. This will preserve the sample.
Although DNA itself is very stable, there might still be some other proteins in the sample that will degrade it over time. The purpose of this protocol is to clean up the saliva sample as much as possible whilst retaining the DNA. Storing the sample in the freezer will slow down any reactions from left over proteins and therefore the template DNA sample will be preserved longer.
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Just as it is vital to clean your kitchen after cooking a meal, laboratory workbenches need to be cleaned between uses. Keeping your lab bench clean ensures leftover dirt or other materials do not contaminate future projects. You can also ensure dangerous substances don’t end up in the wrong places, possibly damaging equipment or cause a colleague harm.
Lab directors should check their departments for cleanliness, but everyone is in charge of keeping work stations clean. A basic check does not need to impact lab processes but should be done every so often.
- Aisles should be clear of boxes, supplies, or other obstructions.
- Loose wires, cables, and computer cords should be tied and organized.
- Lab floors should be mopped at least daily
- Anti-fatigue mats should be replaced regularly to avoid dangerous ware.
- Emergency areas like eyewash stations, showers, and fire extinguishers should remain unobstructed at all times.
- Lab areas should be dusted and de-cluttered regularly.
- Clean items that may not be part of regular testing or experiments. Wipe down chairs, telephones, computers, timers, pens, etc. daily to make sure contagions aren’t accidentally left behind.
Due to the use of chemicals and other hazardous materials used in laboratories, work stations should be more than organized they should be disinfected. Disinfection needs to happen after any spill as well as after every work shift. The CDC recommends the use of a ten percent bleach solution as the standard for disinfection, but other products might be preferred for your lab.
While some manufacturers of laboratory equipment may recommend specific cleaners on their gear, make sure the recommended cleaners are effective for the materials and chemicals that you use. You should also be aware that bleach can cause damage to some lab instruments. Ultimately, you want to use the correct product for the environment in which you are working.
1. Always use the appropriate protective gear. At a minimum, wear latex gloves and goggles. If you have longer hair, tie it up out of the way.
2. Remove loose items from the laboratory work station . Beakers, test tubes, pipettes, etc. should be relocated and washed appropriately.
3. To meet the minimum, 10 percent bleach, mix one-part bleach with nine parts water. This should be sufficient for most lab surfaces. Research your specific station material in case a different solution is required.
4. Dip a paper towel in the mixture and wipe the workbench surface thoroughly. Don’t forget to clean corners, edges, and undersides. You may need to use a wire brush or other device to remove some residue.
- Caked-on material such as solidified agar or other gelatin-like products can be removed by boiling purified water in the equipment.
- Organic materials, including soap residue, can be removed by rinsing with acetone.
- An ethanol rinse is useful to sterilize lab equipment that requires all microorganisms to be removed before use.
- RNAse Displace works well for equipment used in DNA research.
Keeping a clean, disinfected, and tidy workstation will ensure that your experiments and projects are accurate while protecting you and your colleagues from harmful chemicals. It is always better to over-clean than to under-clean. While most surfaces can be cleaned with a bleach solution, check with your specific equipment for requirements and recommendations.
Do you have tips for keeping your lab bench clean? Share them in the comments below.
5 Sure-Fire Ways to Screw Up Your RNA extraction
Working with RNA is definitely an acquired skill. It’s a lot more finicky than working with DNA, and requires careful attention to detail in order to avoid disastrous through RNase contamination. Here are a few common ways to lose your hard-earned RNA:
1. Don’t keep everything on ice
Keeping the temperature of all of your reagents cool is important to inhibit the activity of any luring RNases. If you’re extracting RNA from live cells, it’s okay to spin down the cells at room temperature, but as soon as they’re lysed, you want to keep everything on ice as much as possible. Pre-chilling your buffers can help keep samples cool during manipulation – I’ve even heard of people chilling their pipets and tips in the cold room before working with delicate samples.
2. Don’t Use standard pipet tips
Well-used pipets can get pretty grungy in places you wouldn’t imagine – including the inside of the pipet barrel. Keep this gunk from contaminating your samples by using filter tips, which allow suction from the pipet, but block any particulate matter.
3. Don’t Wear Gloves
Bare hands are the number one source of RNase contamination in the lab. Always use gloves when working with RNA, and remember to change gloves frequently, especially after you’ve touched something else. For kits or reagents that are used exclusively for RNA work, it’s a good idea to put gloves on before even opening the box, to be doubly sure that you’re not introducing any RNases.
4. Use an old/open box of tubes
“Sterile” tubes stored on your bench are often not nearly as sterile as you’d like to think…especially after you’ve dipped into your stash five or ten times. Each time you open the box is another chance to introduce contaminants and RNases. Unless you have exceptional sterile technique, try to use a fresh batch of tubes for any experiment involving RNA.
5. Don’t Use RNase-free Water and Reagents
Most kits for RNA extraction include RNase-free water to use for elution. Handle this bottle of water with gloves as you do with every reagent destined for use with RNA. Don’t be tempted to grab your normal bottle of buffer off of your bench – it’s much better to be safe than sorry. Several companies also offer RNase inhibitor sprays that can be used to clean your bench and gloves before starting a delicate experiment.
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2.1: Using micropipettes correctly
- Contributed by Clare M. O&rsquoConnor
- Associate Professor Emeritus (Biology) at Boston College
Arguably, the most important scientific equipment that you will use in this class are adjustable micropipettes, which you will use in nearly every experiment. Micropipettes are precision instruments that are designed to accuratelyand preciselytransfer volumes in the microliter range. You may use microliters or milliliters as the units of volume in your lab notebooks and lab reports, but be careful to always state the volume unit that you are using. Recall the relationships between volume units:
Accuracy and precision
Accuracy depends on the micropipette delivering the correct volume. Precise results
are reproducible. Let&rsquos use a target analogy to demonstrate the difference between accurate and precise results. Imagine that four students try to hit the bulls-eye five times. Students A and B are precise, while students A and C are accurate.
Manufacturers determine the accuracy and precision of micropipettes by using them to transfer defined volumes of distilled water to a drop that is then weighed on an analytical balance. The density of water is 1.0 gram per mL at 25 ̊C. The process is repeated several times during the calibration process, and the data is used to calculate the accuracy and precision of a micropipette.
Accuracy refers to the performance of the micropipette relative to a standard (the intended) value. Accuracy is computed from the difference between the actual volume dispensed by the micropipette and the intended volume. Note that this can be a negative or positive
value. When micropipettes are calibrated, the accuracy is normally expressed as a percent of
the selected value. Micropipettes are designed to operate with accuracies within a few percent (generally <3%) of the intended value. The accuracy of a micropipette decreases somewhat when micropipettes are set to deliver volumes close to the lowest values in their range.
Precision provides information about reproducibility, without any reference to a standard. Precision reflects random errors that can never be entirely eliminated from a procedure. Thus, a series of repeated measurements should generate a normal or binomial distribution (opposite). Precision is expressed as the standard deviation (s) of the set of measurements. In a normal distribution,
2/3 of measurements will fall within one standard deviation of the average or mean (x), and 95% of measurements will fall within two standard deviations of the mean. The standard deviation for a set of n measurements is calculated using the formula below.
Choosing the micropipette
Standard deviation describes the distribution of measurments relative to the mean value
We use three different sizes of micropipettes in the laboratory, the P20, P200 and P1000. Our micropipettes have been purchased from several different manufacturers, but the principles of operation are the same. The numbers after the &ldquoP&rdquo refer to the maximum number of microli- ters that the micropipette is designed to transfer. Note that there is some overlap in the ranges of the different micropipettes. For example, both the P200 and P20 can be used to transfer 15 &mul, but the P20 is more accurate within that range. As a rule of thumb, always select the smallest volume pipette that will transfer the volume.
Specifying the transfer volume
There are three numbers on the volume indicator. With each of the micropipettes, you will specify a volume to three digits by turning the volume adjustment knob. You will also be able to extrapolate between the lowest numbers with the vernier marks on the lower dial. Most of the measurements you will make with the micropipettes will be accurate to four significant figures!
NEVER turn the indicator dial beyond the upper or lower volume limits of the micropipette! This could damage the piston.
Transferring volumes accurately
Micropipettes work by air displacement. The operator depresses a plunger that moves an internal piston to one of two different positions. The first stop is used to fill the micropipette tip, and the second stop is used to dispense the contents of the tip. As the operator depresses the plunger to the first stop, an internal piston displaces a volume of air equal to the volume shown on the volume indicator dial. The second stop is used only to dispense the contents of the tip.
Filling the micropipette
- Remove the lid from the box containing the correct size micropipette tips. P-1000 tips may be blue or clear, while P-20 and P-200 tips are yellow or clear.
- Attach the tip by inserting the shaft of the micropipette into the tip and pressing down firmly (figure on right). This should produce an airtight seal between the tip and the shaft of the micropipette.
- Replace the lid of the tip box to keep the remaining tips sterile. Avoid touching the tip (especially the thinner end), because the tips are sterile.
- Depress the plunger of the micropipette to the FIRST stop.
- Immerse the tip a few millimeters below the surface of the solution being drawn up into the
pipette. Pipetting is most accurate when the pipette is held vertically. Keep the angle less
than 20 ̊ from vertical for best results.
that the full volume of sample has entered the tip. Do NOT let the plunger snap up. This is particularly important when transferring larger volumes, because a splash could contami- nate the shaft of the micropipette. If you inadvertently contaminate the shaft, clean it imme- diately with a damp Kimwipe.
NEVER rest a micropipette with fluid in its tip on the bench!
Dispensing the contents of the micropipette
- Essay on the Discovery of Enzymes
- Essay on Restriction Endonuclease Enzyme
- Essay on Ligase Enzymes
- Essay on Alkaline Phosphatase Enzyme
- Essay on Phosphonucleotide Kinase Enzyme
- Essay on S1 Nuclease Enzyme
- Essay on DNA Polymerase I: Holoenzyme
- Essay on DNA Polymerase I: Klenow Fragment
- Essay on T4 DNA Polymerase Enzyme
- Essay on Taq DNA Polymerase Enzyme
- Essay on Ribonuclease
- Essay on Deoxyribonuclease I (Dnase I)
- Essay on Terminal Deoxynucleotidyl Transferase Enzyme
- Essay on Reverse Transcriptase Enzyme
Essay # 1. Discovery of Enzymes:
Genetic engineering was born because scientists learned to manipulate DNA. This skill was derived mainly from the field of nucleic acid enzymology. Prior to 1970, there was simply no technique available for cutting a duplex (double-stranded) DNA molecule into distinct fragments. Discovery of DNA metabolising enzymes granted scientists to propose and initiate genetic engineering.
All sort of DNA research developed from the ability, to cut DNA molecules at defined sequences. In other words, it was based upon the discovery of type II restriction endonuclease enzymes.
The isolation of the first restriction endonuclease enzymes, such as Hind II and Hind III, was a result of an interesting discovery by Hamilton O. Smith and his coworkers (1971) that Haemophilus influenza extracts contained activities that cut large DNA molecules into defined fragments.
H. influenza is a non-motile, gram-negative, facultative, anaerobic, pathogenic, rod shaped bacterium which is associated with human respiratory infections, conjunctivitis and meningitis.
Smith’s discovery gave rise to recombinant DNA research. In addition, an entire industry developed with the main purpose of discovery, characterization, purification and marketing of over 100 different site-specific restriction enzymes.
The bringing together of DNA fragments to form covalently linked chimeric molecules is the basis of recombinant DNA research. This step is essential in genetic engineering. This is attained by ligation which is catalysed by DNA ligase, an enzyme which was discovered much prior to that of restriction enzymes.
Before 1970 the existing central dogma in molecular biology was that genetic information transfer occurred from DNA to RNA, and then to protein. The proof that RNA-to-DNA information transfer did occur is based on the discovery and characterization of reverse transcriptase enzyme by Temin and Baltimore (1970).
Reverse transcriptase enzyme allows scientists to generate DNA copies (cDNA) of mRNA subsequent to cloning. The generation of cDNA, containing direct protein coding information is the normal step in cloning of eukaryotic genes.
In fact the most revolutionary and most simplistic molecular biological technical development, is PCR (i.e., polymerase chain reaction). PGR is a direct application of DNA polymerase to permit the test tube binomial amplification of specific DNA sequences.
The discovery of type II restriction enzymes demonstrated the enormous power and utility of site specific DNA cleavage reagents. This article deals with the restriction enzymes and other useful enzymes which are commonly used in genetic engineering (Table 55.1).
Essay # 2. Restriction Endonuclease Enzyme:
Restriction endonucleases (RE) are special class of endonucleases which cleave DNA molecules only at specific nucleotide sequences, called restriction sites. These specific sequences are of four to six nucleotides.
A tetranucleotide sequence will occur more frequently in a given molecule than hexanucleotide, therefore, more fragments will be produced by an enzyme which recognizes tetranucleotide sequence. At these restriction sites, restriction endonucleases cut the DNA by cleaving two phosphodiester bonds one within each strand of the double stranded DNA.
The term ‘restriction endonuclease’ was coined by Lederberg and Meselson (1964) to describe the nuclease enzymes that destroy (‘restrict’) any foreign DNA entering the host cell. However, the first restriction endonuclease enzyme to be isolated and studied was E. coli K12 by Meselson and Yuan (1968). Now these enzymes have been classified into three different types viz., Type I, Type II and Type III.
The discovery of these enzymes led to Nobel Prize for W. Arber, H. Smith and D. Nathans in 1978. In gene manipulation technology, restriction endonuclease enzymes are popularly called molecular knives, molecular scissors or molecular scalpels.
While exact cutting of DNA molecule is very useful for DNA cloning, its full potential is only exhibited when the fragments produced are joined together to give a new structure, known as recombinant DNA. This joining or ligation is achieved by the use of a DNA ligase enzyme.
The most common ligase enzyme is isolated from the bacterial virus (i.e., T4 bacteriophage). Thus, DNA ligases form a group of enzymes which mediate annealing, sealing or joining of DNA fragments. Primarily, ligase enzymes are involved in the repair of DNA molecule where sealing or union of DNA fragments takes place.
DNA ligases also play active part in processes such as DNA replication and recombination. These enzymes are widely used in genetic engineering for the production of hybrid DNA. Since ligase enzymes join DNA fragments or seal the nicks in the chain, they are called molecular structures.
Activity of DNA Ligase Enzymes:
(i) Ligation of DNA molecules with sticky or cohesive ends:
If two different DNA preparations are treated with the same restriction enzyme to give fragments with sticky ends, these ends will be identical in both preparations. Thus, when the two sets of DNA fragments are mixed, base pairing between sticky ends will result in the coming together of fragments which were derived from different molecules.
Also there will be pairing of fragments derived from the same molecule. Such pairing are temporary, owing to the weakness of hydrogen bonding between the few bases in the sticky ends.
The pairing can be stabilised by the use of DNA ligase, which forms a covalent bond between the 5′-phosphoryl at the end of one strand and 3′-hydroxyl of the adjacent strand. Polynucleotide ligase enzyme of T4 bacteriophage catalyzes the end to end joining of DNA duplexes at the base paired end. This reaction could occur intermolecularly or intramolecularly. Researches confirm that intermolecular mode of reaction is correct.
The ligation reaction is driven by ATP and is carried out at 4°C to lower the kinetic energy of molecules. This reduces separation of paired sticky ends and are later stabilised by ligation. However, long reaction time is required to compensate for the low activity of DNA ligase enzyme in the cold. The enzyme concentration is kept high and polyethylene glycol is added to reaction mixture for Stimulation.
Since ligation reconstructs the site of cleavage, recombinant DNA molecules produced by ligation of sticky ends can be cleaved again at the joints, using the same restriction endonuclease enzymes that was used to generate the fragment initially. As a result, a fragment can be inserted into a vector DNA, and recovered again after cloning of the recombinant molecules.
(ii) Ligation of DNA molecules with blunt ends:
Fragments of blunt-ended DNA can be ligated, but since there is no base-pairing to hold fragments together temporarily, concentrations of DNA and ligase enzyme must be high. However, blunt-end ligation is a useful way of joining together DNA fragments which have not been produced by the same restriction enzyme, and which therefore have mismatched sticky ends. These ends are removed prior to ligation, using the enzyme S1 nuclease, which digests single-stranded DNA.
In case of ligation of blunt-ends, a restriction site will not be regenerated and this may prevent recovery of a fragment after cloning. For this reason, short DNA duplexes, called linkers, are frequently used for joining DNA.
Linkers are short, double-stranded oligonucleotides, with blunt ends, containing at least one restriction site (i.e., Eco RI palindrome) within their sequence. These linkers can be joined to one preparation of DNA by blunt- ended ligation and then sticky ends can be created by cleavage of the linkers with a suitable restriction enzyme.
The linker is chosen so that the sticky end it produces is identical to that on the other DNA preparation. Consequently, the two can then be joined by ligation of their sticky ends. Some very versatile linkers are available which contain restriction sites for several different enzymes within a sequence of only eight to ten nucleotides. Blunt- ended molecules can also be ligated after building sticky ends.
Thus, with this method it is now possible to insert a foreign DNA segment at a particular site in the linker region of the vector and then retrieve this foreign DNA segment whenever necessary.
Sources of DNA Ligases:
DNA ligases are isolated from E. coli and T4 bacteriophage. The ligase enzyme isolated from E. coli is a polypeptide chain with a molecular weight of 75 kDa. It requires NAD + as cofactor. The ligase obtained from T4 bacteriophage is 68 kDa. It requires ATP as a cofactor and a source of energy.
For the routine laboratory requirement, T4 DNA ligase is obtained from an induced lysogen of lambda T4 lig phage. This enzyme has the capacity to ligate a variety of cohesive and blunt- ended DNA fragments. The enzyme concentration is kept higher and a fusogen called polyethylene glycol, is added to reaction mixture for stimulation.
Application of DNA Ligase Enzymes:
DNA ligase enzymes play an important role in genetic engineering. In the absence of DNA ligase enzymes, recombinant DNA technology cannot be successful.
The important functions of DNA ligase enzymes are as follows:
1. Genetic engineering experiments involve joining of DNA fragments to produce recombinant DNA molecules. Ligase enzymes are used in the joining process.
2. Ligase enzymes help in ligation of vector and inserting recombinant DNA.
3. They help in ligation of linkers or adapter molecules at the blunt ends of DNA fragments.
4. They help in sealing nicks in double- stranded DNA.
5. Ligase enzymes requires 3′ OH and 5′ P0^4 group for ligation.
This requirement can be advantageous as:
(i) Self ligation of DNA can be prevented by dephosphorylation (using alkaline phosphatase) of intended donor fragments prior to ligation.
(ii) Dephosphorylation of vector DNA will prevent recircularization of the vector in cloning procedure.
Essay # 4. Alkaline Phosphatase Enzyme:
The enzyme alkaline phosphatase (AP) catalyses the removal of the 5′-terminal phosphate residues from nucleic acids (RNA, DNA and ribo- and deoxyribonucleotide triphosphates). This enzyme is isolated from bacteria (BAP) or calf intestine (CAP).
This enzyme is a dimeric glycoprotein with a molecular weight 14,000. It is made up of two identical or similar subunits each with a molecular weight of 6900. It is a zinc-containing enzyme with four atoms of Zn 2+ per molecule.
Uses of Alkaline Phosphatase Enzyme:
1. Linearized cloning vectors can be prevented from recircularizing by dephosphorylation with alkaline phosphatase enzyme.
2. The free 5′-OH can be phosphorylated with polynucleotide kinase and ϒ -32P ATP to produce 32P end labelled nucleic acid.
3. AP enzyme is used for mapping and DNA fingerprinting studies.
Essay # 5. Phosphonucleotide Kinase Enzyme:
The enzyme phosphonucleotide kinase catalyses the transfer of the terminal phosphate group of ATP to the 5′-hydroxylated terminal of DNA or RNA. This enzyme is frequently used to end- label the nucleic acids with 32 P (i.e., it adds the phosphate back to 5′-termini of DNA).
This can be accomplished by any method among following:
1. Forward reaction:
Transfer of labelled ϒ-phosphate form (ϒ -32 P)-ATP to the free 5′-hydroxyl group of substrate-
5′ – OH – DNA + [ -32 P] ATP5′ 32 → PO – DNA + ADP
Substrate lacking a free 5′-hydroxyl requires prior dephosphorylation by alkaline phosphatase.
2. Exchange reaction:
In the initial step, the terminal 5′-phosphate is transferred from substrate to ADP present in the reaction mixture. Then, the labelled ϒ-phosphate from [ϒ- 32 P]-ATP is transferred to free hydroxyl group of substrate.
5′ – HO – DNA + [ϒ -32 P] – ATP → 5′ 32 PO – DNA + ATP
Uses of Polynucleotide Kinase Enzyme:
The enzyme polynucleotide kinase is used to label 5′-termini of DNA and RNA with [ϒ- 32 P]-ATP by phosphorylation of 5′-hydroxyl groups or by the exchange reaction. This 5′-terminal labelling is used in mapping of restriction sites, DNA or RNA fingerprinting, hybridization studies and sequence analysis of DNA.
Essay # 6. S1 Nuclease Enzyme:
The S1 nuclease enzyme is single- strand specific endonuclease which cleaves DNA to release 5′-mono and 5′-oligonucleotides. Normally, double- stranded DNA, double- stranded RNA and DNA-RNA hybrids are resistant to action of S1 nuclease enzyme.
However, very large amounts of S1 nuclease enzyme can completely hydrolyze double- stranded nucleic acids. The enzyme hydrolyzes single stranded regions in duplex DNA such as loops and gaps.
S1 nuclease enzyme can also cleave single stranded areas of super helical DNA at torsional stress points where DNA may be unpaired or weakly hydrogen bonded. Once the super-helical DNA is nicked, S1 nuclease enzyme can cleave the second strand near the nick to generate linear DNA.
S1 nuclease enzyme is a monomeric protein with 3800 dalton molecular weight. It requires Zn 2+ for its activity and is relatively stable against denaturing reagents such as urea, SDS and formamide. The optimum pH requirement lies between 4 to 4.5.
Uses of S1 Nuclease Enzyme:
1. S1 nuclease enzyme is used to analyse DNA-RNA hybrid structures to map transcripts.
2. It can be used to remove singles stranded tails from DNA fragments to produce blunt ends.
3. Hair pin loop structures formed during synthesis of double-stranded cDNA is digested by this enzyme.
4. S1 nuclease enzyme is also used for DNA mapping, called SI nuclease mapping Turner.
Essay # 7. DNA Polymerase I: Holoenzyme:
This enzyme has two-fold activities:
5′ → 3’exonuclease activity and DNA synthesis (acts as 5’ -3’ polymerase). Such a bifunctional activity enables the DNA polymerase I enzyme to use nicks or gaps in double- a stranded DNA as a starting point of DNA synthesis.
The 5′-exonuclease activity degrades that DNA strand which is complementary to the template strand and thus forming nick. DNA synthesis begins at 3′-end of nick and produces a new strand of DNA complementary to the template.
The new result is the movement of nick along the template strand (nick translation) until all the DNA complementary to the template strand (starting from the site of the origin of nick to the 5′-end of the template strand) is replaced.
Uses of DNA Polymerase I:
1. DNA polymerase I enzyme is used with radioactive or biotinylated nucleotides to prepare labelled DNA of high specific activity.
2. DNA polymerase I enzyme can also catalyse de novo DNA synthesis.
3. The enzyme DNA polymerase I has 3’→5′ proof-reading exonuclease activities on a single polynucleotide chain.
Essay # 8. DNA Polymerase I: Klenow Fragment:
Treatment of DNA polymerase I holoenzyme of E. coli with protease enzyme results in the production of two protein fragments. The larger fragment is called Klenow fragment and it does not show 5′-exonuclease activity (i.e., the 5′- exonuclease activity is exhibited by the intact enzyme) This Klenow fragment is used to synthesize DNA when there is no need of removing the DNA strand which is complementary to the template strand.
Uses of Klenow Fragment:
Klenow fragments are used in the following ways:
1. In DNA sequencing by dideoxy method.
2. For the production of second strand of cDNA.
3. Radiolabelling by filling in 5′-single stranded extension on double-stranded DNA.
4. Mutagenesis of DNA with synthetic oligonucleotides.
5. In labelling the DNA by random primer method.
+ stands for presence of particular activity by the enzyme.
0 presents absence of such activity.
Essay # 9. T4 DNA Polymerase Enzyme:
The enzyme T4 DNA polymerase lacks 5′-exonuclease activity but has a very active 3′- exonuclease action. This property allows radiolabelling of DNA fragment by replacement synthesis.
In method of replacement synthesis, T4 DNA polymerase enzyme acts on both the 3’ -ends of double sanded extensions. DNA complementary to these single strands are synthesized with deoxynucleoside triphosphate enzyme in the presence of radiolabelling compounds.
Application of T4 DNA polymerase:
1. T4 DNA polymerase is useful in generating 5′ single-stranded ends.
2. The enzyme can be used in radiolabelling of DNA.
Essay # 10. Taq DNA Polymerase Enzyme:
The enzyme Taq to DNA polymerase is isolated from the thermophilic bacterium Thermus aquaticus. Taq enzyme has the highest DNA polymerase activity at a pH of 9 and temperature around 75°C.
A Activity of Taq DNA polymerase is resistant to incubation at high as 95°C. Taq enzyme consists of a single polypeptide chain with a molecular weight of 95000. It lacks 5′ to 3′ and 3′ to 5′ exonuclease activity.
The highly thermostable Taq DNA polymerase from Thermus aquaticus is ideal for both manual and automated DNA sequencing because it is fast, highly progressive, has little or no 3’ – exonuclease activity and is active over a broad range of temperature.
Application of Taq Polymerase:
1. Taq enzyme is used in DNA sequencing studies.
2. Taq enzyme is used in ‘Polymerase chain reaction’ or PGR as it can withstand high temperatures.
Essay # 11. Ribonuclease:
Generally RNase A and RNase T1 enzymes are used in genetic engineering techniques. Both enzymes cleave the phosphodiester bond between adjacent ribonucleotides. RNase A cleaves next to uracil (U) and cytosine (C) in such a way that phosphate remains with these pyrimidines. The nucleotide present on the other side of phosphate is dephosphorylated. RNase A enzyme is isolated from the bovine pancreas.
RNase T1 cleaves specifically next to guanine. The phosphate group at the 3′ end of the nucleotide remains with the cut end. This enzyme is isolated from Aspergillus oryzae.
Ribonuclease H (RNase H):
The enzyme RNase H is an endoribonuclease that degrades the RNA portion of the RNA- DNA hybrids. RNase H enzyme cuts the RNA into short fragments.
Applications of RNase H:
1. RNase H is the key enzyme in the cDNA cloning technique. In this case, it is used to remove the mRNA from the RNA-DNA hybrid.
2. RNase H enzyme is used to detect the presence of RNA-DNA hybrid.
3. RNase H enzyme is used to remove poly (A) tails on mRNA.
The enzyme poIy-A polymerase plays a vital role in vitro gene manipulation techniques as it analyses the addition of AMP units to the 3′ end of RNA.
Essay # 12. Deoxyribonuclease I (Dnase I):
The enzyme DNase I is an endonuclease enzyme which digests either single or double-stranded DNA, producing a mixture of mononucleotides and oligonucleotides. DNase I hydrolyses each strand of double-stranded DNA independently and at random. Addition of Mg 2+ to reaction mixture ensures random cleavage while addition of Mn 2+ gives cleavage nearly at the same place on both strands. DNase enzyme is obtained mostly from bovine pancreas.
Uses of DNase I Enzyme:
DNase 1 enzyme is useful for a variety of applications including nick translation, DNA foot printing, bisulphite mediated mutagenesis and RNA purification.
Essay # 13. Terminal Deoxynucleotidyl Transferase Enzyme:
The enzyme deoxynucleotidyl transferase catalyses the repetitive addition of monodeoxynucleotide units from a deoxynucleoside triphosphate to the terminal 3′-hydroxyl group of a DNA molecule. This enzyme has a molecular weight of 32000 and consists of two subunits each with a molecular weight of 26500 and 8000. This enzyme is isolated from calf thymus.
Uses of Terminal Transferase Enzyme:
1. The enzyme terminal transferase is used to add homopolymer tails of DNA fragmeirts. Using a technique called homopolymer tailing, sticky ends can be built up on blunt-ended DNA molecules.
For examples, one preparation of DNA could be treated with the enzyme terminal transferase in the presence of dATP, resulting in the addition of a poly (dA) chain to each DNA strand. There is another preparation of DNA which provides 3 tails of poly (T) using same enzyme with TTP.
When both types of DNA preparations DNA fragments with poly A tails and DNA fragments with poly T tails, are mixed, there takes place base pairing between complementary sticky ends, which could then be ligated. One advantage of this method is that ligation does not take place between fragments from the same DNA preparation.
2. Terminal transferase enzyme is used for 3′-end labelling of DNA fragments
3. Terminal transferase enzyme is also used for the addition of single nucleotides to the 3- end of DNA for in vitro mutagenesis.
Purine and pyrimidine polymerization rates, by using terminal transferase enzyme, depend on the addition of Mg 2+ , Mn 2+ or Co 2+ in the reaction mixture.
Essay # 14. Reverse Transcriptase Enzyme:
The enzyme reverse transcriptase is isolated from avian myeloblastos virus (AMV). It is an RNA-dependent DNA polymerase. The enzyme requires DNA primer complementary to the RNA template, as well as presence of Mg 2+ or Mn 2+ for initiation of transcription. Reverse transcriptase mediates the conversion of genetic information present in single- stranded molecule of RNA into a double-stranded molecule of DNA.
Until recently, it was known that the genetic information’s of DNA pass to protein through mRNA. During 1960s, Temin and coworkers postulated that in certain cancer causing animal viruses which contain RNA as genetic material, transcription of cancerous genes (on RNA into DNA) takes places most probably by DNA polymerase directed by viral RNA.
Then DNA is used as template for synthesis of many copies of viral RNA in a cell. In 1970, S. Mizutani, H.M. Temin and D. Baltimore discovered that information can pass back from RNA to DNA.
They found that retroviruses (possessing RNA) contain RNA dependent DNA polymerase which IS also called reverse transcriptase. This process produces single- stranded DNA which in turn functions as template for complementary chain of DNA.
Reverse transcriptase enzyme has two subunits. The enzymatically active forms of the purified enzyme are α, β and αβ. The molecular weight of the α-subunit is 68000 and that of β-subunit is 92,000.
The mature α-β form is the most active form of AMV reverse transcriptase enzyme. It has several enzymatic roles such as RNA-directed DNA polymerase action, DNA dependent RNA polymerase activity and RNase-H activity.
The α- subunit of reverse transcriptase contains the polymerase activity. It also has the RNase- H activity during which degradation of RNA in DNA: RNA hybrids takes place. Such a sort of exonucleolytic activity of RNase-H enzyme can proceed either from the 5′- or 3′- terminus.
Uses of Reverse Transcriptase Enzyme:
1. The in vitro synthesis of cDNA from mRNA and other RNA molecule using reverse transcriptase enzyme has become a very important technique in the field of molecular biology.
2. DNA-dependent DNA polymerase activity of reverse transcriptase enzyme is responsible for second-strand formation in cDNA synthesis. Such a polymerising activity of reverse transcriptase is inhibited by the addition of actinomycin-D.
3. The reverse transcriptase enzyme mediates the conversion of genetic information present in single-stranded molecule of RNA into a double-stranded molecule of DNA.
Ribonuclease H is a family of endonuclease enzymes with a shared substrate specificity for the RNA strand of RNA-DNA duplexes. By definition, RNases H cleave RNA backbone phosphodiester bonds to leave a 3' hydroxyl and a 5' phosphate group.  RNases H have been proposed as members of an evolutionarily related superfamily encompassing other nucleases and nucleic acid processing enzymes such as retroviral integrases, DNA transposases, Holliday junction resolvases, Piwi and Argonaute proteins, various exonucleases, and the spliceosomal protein Prp8.  
RNases H can be broadly divided into two subtypes, H1 and H2, which for historical reasons are given Arabic numeral designations in eukaryotes and Roman numeral designations in prokaryotes. Thus the Escherichia coli RNase HI is a homolog of the Homo sapiens RNase H1.   In E. coli and many other prokaryotes, the rnhA gene encodes HI and the rnhB gene encodes HII. A third related class, called HIII, occurs in a few bacteria and archaea it is closely related to prokaryotic HII enzymes. 
The structure of RNase H commonly consists of a 5-stranded β-sheet surrounded by a distribution of α-helices.  All RNases H have an active site centered on a conserved sequence motif composed of aspartate and glutamate residues, often referred to as the DEDD motif. These residues interact with catalytically required magnesium ions.  
RNases H2 are larger than H1 and usually have additional helices. The domain organization of the enzymes varies some prokaryotic and most eukaryotic members of the H1 group have an additional small domain at the N-terminus known as the "hybrid binding domain", which facilitates binding to RNA:DNA hybrid duplexes and sometimes confers increased processivity.    While all members of the H1 group and the prokaryotic members of the H2 group function as monomers, eukaryotic H2 enzymes are obligate heterotrimers.   Prokaryotic HIII enzymes are members of the broader H2 group and share most structural features with H2, with the addition of an N-terminal TATA box binding domain.  Retroviral RNase H domains occurring in multidomain reverse transcriptase proteins have structures closely resembling the H1 group. 
RNases H1 have been extensively studied to explore the relationships between structure and enzymatic activity. They are also used, especially the E. coli homolog, as model systems to study protein folding.    Within the H1 group, a relationship has been identified between higher substrate-binding affinity and the presence of structural elements consisting of a helix and flexible loop providing a larger and more basic substrate-binding surface. The C-helix has a scattered taxonomic distribution it is present in the E. coli and human RNase H1 homologs and absent in the HIV RNase H domain, but examples of retroviral domains with C-helices do exist.  
Ribonuclease H enzymes cleave the phosphodiester bonds of RNA in a double-stranded RNA:DNA hybrid, leaving a 3' hydroxyl and a 5' phosphate group on either end of the cut site with a two-metal-ion catalysis mechanism, in which two divalent cations, such as Mg2+ and Mn2+, directly participate in the catalytic function.  Depending on the differences in their amino acid sequences, these RNases H are classified into type 1 and type 2 RNases H.   Type 1 RNases H have prokaryotic and eukaryotic RNases H1 and retroviral RNase H. Type 2 RNases H have prokaryotic and eukaryotic RNases H2 and bacterial RNase H3. These RNases H exist in a monomeric form , except for eukaryotic RNases H2, which exist in a heterotrimeric form.   RNase H1 and H2 have distinct substrate preferences and distinct but overlapping functions in the cell. In prokaryotes and lower eukaryotes, neither enzyme is essential, whereas both are believed to be essential in higher eukaryotes.  The combined activity of both H1 and H2 enzymes is associated with maintenance of genome stability due to the enzymes' degradation of the RNA component of R-loops.  
Ribonuclease H1 Edit
Ribonuclease H1 enzymes require at least four ribonucleotide-containing base pairs in a substrate and cannot remove a single ribonucleotide from a strand that is otherwise composed of deoxyribonucleotides. For this reason, it is considered unlikely that RNase H1 enzymes are involved in the processing of RNA primers from Okazaki fragments during DNA replication.  RNase H1 is not essential in unicellular organisms where it has been investigated in E. coli, RNase H1 knockouts confer a temperature-sensitive phenotype,  and in S. cerevisiae, they produce defects in stress response. 
In many eukaryotes, including mammals, RNase H1 genes include a mitochondrial targeting sequence, leading to expression of isoforms with and without the MTS present. As a result, RNase H1 is localized to both mitochondria and the nucleus. In knockout mouse models, RNase H1-null mutants are lethal during embryogenesis due to defects in replicating mitochondrial DNA.    The defects in mitochondrial DNA replication induced by loss of RNase H1 are likely due to defects in R-loop processing. 
Ribonuclease H2 Edit
In prokaryotes, RNase H2 is enzymatically active as a monomeric protein. In eukaryotes, it is an obligate heterotrimer composed of a catalytic subunit A and structural subunits B and C. While the A subunit is closely homologous to the prokaryotic RNase H2, the B and C subunits have no apparent homologs in prokaryotes and are poorly conserved at the sequence level even among eukaryotes.   The B subunit mediates protein-protein interactions between the H2 complex and PCNA, which localizes H2 to replication foci. 
Both prokaryotic and eukaryotic H2 enzymes can cleave single ribonucleotides in a strand.  however, they have slightly different cleavage patterns and substrate preferences: prokaryotic enzymes have lower processivity and hydrolyze successive ribonucleotides more efficiently than ribonucleotides with a 5' deoxyribonucleotide, while eukaryotic enzymes are more processive and hydrolyze both types of substrate with similar efficiency.   The substrate specificity of RNase H2 gives it a role in ribonucleotide excision repair, removing misincorporated ribonucleotides from DNA, in addition to R-loop processing.    Although both H1 and H2 are present in the mammalian cell nucleus, H2 is the dominant source of RNase H activity there and is important for maintaining genome stability. 
Some prokaryotes possess an additional H2-type gene designated RNase HIII in the Roman-numeral nomenclature used for the prokaryotic genes. HIII proteins are more closely related to the H2 group by sequence identity and structural similarity, but have substrate preferences that more closely resemble H1.   Unlike HI and HII, which are both widely distributed among prokaryotes, HIII is found in only a few organisms with a scattered taxonomic distribution it is somewhat more common in archaea and is rarely or never found in the same prokaryotic genome as HI. 
The active site of nearly all RNases H contains four negatively charged amino acid residues, known as the DEDD motif often a histidine e.g in HIV-1, human or E. coli is also present.  
The charged residues bind two metal ions that are required for catalysis under physiological conditions these are magnesium ions, but manganese also usually supports enzymatic activity,   while calcium or high concentration of Mg2+ inhibits activity.   
Based on experimental evidence and computer simulations the enzyme activates a water molecule bound to one of the metal ions with the conserved histidine.   The transition state is associative in nature  and forms an intermediate with protonated phosphate and deprotonated alkoxide leaving group.  The leaving group is protonated via the glutamate which has an elevated pKa and is likely to be protonated. The mechanism is similar to RNase T and the RuvC subunit in the Cas9 enzyme which both also use a histidine and a two-metal ion mechanism.
The mechanism of the release of the cleaved product is still unresolved. Experimental evidence from time-resolved crystallography and similar nucleases points to a role of a third ion in the reaction recruited to the active site.  
The human genome contains four genes encoding RNase H:
- , an example of the H1 (monomeric) subtype , the catalytic subunit of the trimeric H2 complex , a structural subunit of the trimeric H2 complex , a structural subunit of the trimeric H2 complex
In addition, genetic material of retroviral origin appears frequently in the genome, reflecting integration of the genomes of human endogenous retroviruses. Such integration events result in the presence of genes encoding retroviral reverse transcriptase, which includes an RNase H domain. An example is ERVK6.  Long terminal repeat (LTR) and non-long terminal repeat (non-LTR) retrotransposons are also common in the genome and often include their own RNase H domains, with a complex evolutionary history.   
Role in disease Edit
In small studies, mutations in human RNase H1 have been associated with chronic progressive external ophthalmoplegia, a common feature of mitochondrial disease. 
Mutations in any of the three RNase H2 subunits are well-established as causes of a rare genetic disorder known as Aicardi–Goutières syndrome (AGS),  which manifests as neurological and dermatological symptoms at an early age.  The symptoms of AGS closely resemble those of congenital viral infection and are associated with inappropriate upregulation of type I interferon. AGS can also be caused by mutations in other genes: TREX1, SAMHD1, ADAR, and MDA5/IFIH1, all of which are involved in nucleic acid processing.  Characterization of mutational distribution in an AGS patient population found 5% of all AGS mutations in RNASEH2A, 36% in 2B, and 12% in 2C.  Mutations in 2B have been associated with somewhat milder neurological impairment  and with an absence of interferon-induced gene upregulation that can be detected in patients with other AGS-associated genotypes. 
Two groups of viruses use reverse transcription as part of their life cycles: retroviruses, which encode their genomes in single-stranded RNA and replicate through a double-stranded DNA intermediate and dsDNA-RT viruses, which replicate their double-stranded DNA genomes through an RNA "pregenome" intermediate. Pathogenic examples include human immunodeficiency virus and hepatitis B virus, respectively. Both encode large multifunctional reverse transcriptase (RT) proteins containing RNase H domains.  
Retroviral RT proteins from HIV-1 and murine leukemia virus are the best-studied members of the family.   Retroviral RT is responsible for converting the virus' single-stranded RNA genome into double-stranded DNA. This process requires three steps: first, RNA-dependent DNA polymerase activity produces minus-strand DNA from the plus-strand RNA template, generating an RNA:DNA hybrid intermediate second, the RNA strand is destroyed and third, DNA-dependent DNA polymerase activity synthesizes plus-strand DNA, generating double-stranded DNA as the final product. The second step of this process is carried out by an RNase H domain located at the C-terminus of the RT protein.    
RNase H performs three types of cleaving actions: non-specific degradation of the plus-strand RNA genome, specific removal of the minus-strand tRNA primer, and removal of the plus-strand purine-rich polypurine tract (PPT) primer.  RNase H plays a role in the priming of the plus-strand, but not in the conventional method of synthesizing a new primer sequence. Rather RNase H creates a "primer" from the PPT that is resistant to RNase H cleavage. By removing all bases but the PPT, the PPT is used as a marker for the end of the U3 region of its long terminal repeat. 
Because RNase H activity is required for viral proliferation, this domain has been considered a drug target for the development of antiretroviral drugs used in the treatment of HIV/AIDS and other conditions caused by retroviruses. Inhibitors of retroviral RNase H of several different chemotypes have been identified, many of which have a mechanism of action based on chelation of the active-site cations.  Reverse-transcriptase inhibitors that specifically inhibit the polymerase function of RT are in widespread clinical use, but not inhibitors of the RNase H function it is the only enzymatic function encoded by HIV that is not yet targeted by drugs in clinical use.  
RNases H are widely distributed and occur in all domains of life. The family belongs to a larger superfamily of nuclease enzymes   and is considered to be evolutionarily ancient.  In prokaryotic genomes, multiple RNase H genes are often present, but there is little correlation between occurrence of HI, HII, and HIII genes and overall phylogenetic relationships, suggesting that horizontal gene transfer may have played a role in establishing the distribution of these enzymes. RNase HI and HIII rarely or never appear in the same prokaryotic genome. When an organism's genome contains more than one RNase H gene, they sometimes have significant differences in activity level. These observations have been suggested to reflect an evolutionary pattern that minimizes functional redundancy among RNase H genes.   RNase HIII, which is unique to prokaryotes, has a scattered taxonomic distribution and is found in both bacteria and archaea  it is believed to have diverged from HII fairly early. 
The evolutionary trajectory of RNase H2 in eukaryotes, especially the mechanism by which eukaryotic homologs became obligate heterotrimers, is unclear the B and C subunits have no apparent homologs in prokaryotes.  
Because RNase H specifically degrades only the RNA in double-stranded RNA:DNA hybrids, it is commonly used as a laboratory reagent in molecular biology. Purified preparations of E. coli RNase HI and HII are commercially available. RNase HI is often used to destroy the RNA template after first-strand complementary DNA (cDNA) synthesis by reverse transcription. It can also be used to cleave specific RNA sequences in the presence of short complementary segments of DNA.  Highly sensitive techniques such as surface plasmon resonance can be used for detection.   RNase HII can be used to degrade the RNA primer component of an Okazaki fragment or to introduce single-stranded nicks at positions containing a ribonucleotide.  A variant of hot start PCR, known as RNase H-dependent PCR or rhPCR, has been described using a thermostable RNase HII from the hyperthermophilic archaeon Pyrococcus abyssi.  Of note, the ribonuclease inhibitor protein commonly used as a reagent is not effective at inhibiting the activity of either HI or HII. 
Ribonucleases H were first discovered in the laboratory of Peter Hausen when researchers found RNA:DNA hybrid endonuclease activity in calf thymus in 1969 and gave it the name "ribonuclease H" to designate its hybrid specificity.    RNase H activity was subsequently discovered in E. coli  and in a sample of oncoviruses with RNA genomes during early studies of viral reverse transcription.   It later became clear that calf thymus extract contained more than one protein with RNase H activity  and that E. coli contained two RNase H genes.   Originally, the enzyme now known as RNase H2 in eukaryotes was designated H1 and vice versa, but the names of the eukaryotic enzymes were switched to match those in E. coli to facilitate comparative analysis, yielding the modern nomenclature in which the prokaryotic enzymes are designated with Roman numerals and the eukaryotic enzymes with Arabic numerals.     The prokaryotic RNase HIII, reported in 1999, was the last RNase H subtype to be identified. 
Characterizing eukaryotic RNase H2 was historically a challenge, in part due to its low abundance.  Careful efforts at purification of the enzyme suggested that, unlike the E. coli RNase H2, the eukaryotic enzyme had multiple subunits.  The S. cerevisiae homolog of the E. coli protein (that is, the H2A subunit) was easily identifiable by bioinformatics when the yeast genome was sequenced,  but the corresponding protein was found not to have enzymatic activity in isolation.   Eventually, the yeast B and C subunits were isolated by co-purification and found to be required for enzymatic activity.  However, the yeast B and C subunits have very low sequence identity to their homologs in other organisms, and the corresponding human proteins were conclusively identified only after mutations in all three were found to cause Aicardi–Goutières syndrome.  
Ribonucleases - Part A
Walt F. Lima , . Stanley T. Crooke , in Methods in Enzymology , 2001
RNase H hydrolyzes RNA in RNA–DNA hybrids. 1 RNase H activity appears to be ubiquitous in eukaryotes and bacteria. 2–7 Although RNases H constitute a family of proteins of varying molecular weight, the nucleolytic activity and substrate requirements appear to be similar for the various isotypes. For example, all RNases H studied to date function as endonucleases exhibiting limited sequence specificity and requiring divalent cations (e.g., Mg 2+ , Mn 2+ ) to produce cleavage products with 5′-phosphate and 3′-hydroxyl termini. 8
Two classes of RNase H enzymes have been identified in mammalian cells. 5,9,10 These enzymes were shown to differ with respect to cofactor requirements. For example, RNase H1 is activated by both Mg 2+ and Mn 2+ , whereas RNase H2 was shown to be activated by only Mg 2+ and inhibited by Mn 2+ . Furthermore, both RNase H1 and H2 were shown to be inhibited by sulfhydryl reagents. 10,11 Although the biological roles of the mammalian enzymes are not fully understood, it has been suggested that mammalian RNase H1 may be involved in replication and that the type 2 enzyme may be involved in transcription. 12,13
Recently, both human RNase H genes have been cloned and expressed. 11,14,15 The type 1 enzyme is a 286 amino acid protein with a calculated mass of 32 kDa. 11 The enzyme is encoded by a single gene that is at least 10 kb in length and is expressed ubiquitously in human cells and tissues. The amino acid sequence of human RNase H1 displays strong homology with RNase H1 from yeast (21.8% amino acid identity), chicken (59%), Escherichia coli (33.6%), and mouse (84.3%). The type 1 enzymes are all small proteins (<40 kDa) and their estimated pI values are all 8.7 and greater. The amino acid residues in E. coli RNase HI thought to be involved in the Mg 2+ binding site, catalytic center, and substrate binding region 16–18 are well conserved in the cloned human RNase H1 sequence. 11 The human RNase H2 enzyme is a 299 amino acid protein with a calculated mass of 33.4 kDa and has also been shown to be ubiquitously expressed in human cells and tissues (see Ref. 14 Wu, unpublished data, 1998). Human RNase H2 shares strong amino acid sequence homology with RNase H2 from Caenorhabditis elegans (45.5% amino acid identity), yeast (25.7%), and E. coli (14.4%). Unlike the RNase H1 isotype, the type 2 enzyme is an acidic protein exhibiting a pI of 4.94.