PRACTICAL MANUAL
Molecular Biology
Course Instructor
Dr. Pakeeza A. Shaiq
Teacher’s Assistant
Ms. Sadaf Naz
Mr. Muhammad Mazhar
University Institute of Biochemistry and Biotechnology
Practical#1
Laboratory Ethics
General Rules and Regulations:
It is necessary for everyone working in the lab to observe safety rules and regulations of each laboratory.
Below are general safety guidelines applicable to most laboratories:
1.
Personal Protective Equipment (PPE):

Lab coats or disposable aprons should be worn in the lab to protect you and your clothing from
contamination. Lab coats should not be worn outside the laboratory.

Lab footwear should consist of normal closed shoes to protect all areas of the foot from possible
puncture from sharp objects or broken glass and from contamination from corrosive reagents or infectious
materials.

Gloves should be worn for handling chemicals, blood and specimens.

Protective eyewear such as goggles and masks may need to be worn when contact with hazardous
aerosols, caustic chemicals or reagents.
2.
Eating, drinking, smoking, applying cosmetics and handling contact lenses are prohibited in areas
where specimens are handled.
3.
Smoking is prohibited in all laboratory areas.
4.
Food and drink should not be stored in refrigerators, freezers, cabinets or on shelves where
chemicals, blood or other potentially infectious materials are stored.
5.
Long hair, ties, scarves and earrings should be secured.
6.
Never mouth pipette, mechanical pipetting devices must be used for pipetting all liquids.
7.
Frequent hand washing is an important safety precaution, which should be practiced before and after
every experiment.
8.
Label all storage areas appropriately and keep all chemicals in properly labelled containers.
9.
Note expiry dates on chemicals.
10.
Always return reagents to their proper place in the laboratory immediately after use.
11.
Always return the cap to the proper bottle and ensure that it is secured on the bottle.
12.
Never return unused chemicals to a reagent bottle. Doing so might result in the contamination of the
reagent.
13.
When heating or carrying out reactions in a test tube, never point the mouth of the tube at your
neighbor or yourself.
14.
Never taste a chemical; never smell a chemical unless instructed to do so. If instructed to smell a
chemical, fan vapors toward your nose and inhale cautiously.
15.
Work in a fume hood when handling toxic and fume producing chemicals.
16.
Handle and store glassware carefully so as not to damage it or yourself.
17.
When inserting glass tubing into rubber stoppers, corks or when placing rubber tubing on glass hose
connections protect hands with a heavy glove and lubricate tubing or stopper.
18.
When dealing with broken glass wear hand protection when picking up the pieces. Use a broom to
sweep small pieces into a dustpan and store glass pieces in a designated bin for broken glass.
19.
Paper, broken glass, stoppers, rubber tubing, etc., are to be kept out of sinks at all times to minimize
the possibility of clogged drains. Such items are to be kept away from areas where they might fall into sinks
or drains.
20.
Do not work with flammable solvents/chemicals near open flames (e.g. a gasburner) at the same time
in the same laboratory.
21.
Warning signs should be posted to alert attentions in the work area when unusual hazards, such as
radiation, laser operations, dangerous chemicals, biological hazards, or other special hazards exist.
22.
Before leaving the laboratory electrical equipments should be turned off and gas burners
extinguished. No tap water should be left running.
Hazard Symbols
Biosafety Cabinets
Introduction:
Biosafety cabinets also known as biological safety cabinets (BSCs) are an enclosed, ventilated hood or
workspace that allow for the safe handling of pathogens, contaminants or other potentially hazardous
materials. The primary purpose of a biosafety cabinet is to protect the operator and the surrounding
environment from biological contaminants and other hazardous materials. These cabinets are designed to
provide various levels of protection. There are various classes of biological safety cabinets, each defined by
the required level of biosafety and containment.
Classes of biosafety cabinets:
Class Ι:
The Class I biosafety cabinet provides personnel and environmental protection, but no product
protection. It has a HEPA filter in the exhaust system to provide containment and environmental
protection. This older class of biosafety cabinet is rarely used.
Class ΙΙ:
The Class II biosafety cabinet provides protection to the user, the experimental material and the
environment. Air flow is drawn from the room around the operator into the front grille of the cabinet, which
provides personnel protection. In addition, the downward laminar flow of HEPA-filtered air provides
protection for experimental material inside the cabinet. Because cabinet air has passed through the exhaust
HEPA filter, it is contaminant-free, providing environmental protection, and may be recirculated back into
the laboratory. (Class II Type A) or ducted out of the building (Class II Type B).
Class ΙΙΙ:
The Class III biological safety cabinet is most suitable for work with bio-hazardous agents requiring
high contain (biosafety level 3 or 4). The Class III cabinet is completely enclosed, HEPA filter-ventilated
cabinet fitted with glove ports and decontamination capabilities for entry and exit of material. It offers the
highest degree of personnel and environmental protection from infectious aerosols.
Clean bench & fume hood:
Clean Benches and Fume Hoods are Not Biological Safety Cabinets. Fume hoods are ventilated
enclosures that remove hazardous chemical fumes and volatile vapors from the laboratory, providing
personnel protection only, hence should not be used for biohazard substances. Whereas, a clean bench
provides a space to work with a product or specimen where it will be protected from contamination by
particulates such as microorganisms. This is accomplished by the laminar flow of clean air from a HEPA
filter, which is blown across the workspace and out toward the user and the lab. Thus, clean benches should
not be used when working with potentially infectious materials, chemical hazards or radioactivity.
Biosafety Levels
Definition:
A biosafety level (BSL) is a set of bio-containment precautions required to isolate dangerous
biological agents in an enclosed laboratory facility.
Biosafety levels:
Biosafety Levels 1-4 are combinations of laboratory practices and techniques, safety equipment and
facilities. All of these levels are appropriate for the biohazard posed by the agents used in researches and
experiments.
Biosafety level 1:
Biosafety level one, the lowest level, applies to work with agents that usually pose a minimal
potential threat to laboratory workers and the environment and do not consistently cause disease in healthy
adults. An example of a microbe that is typically worked with at a BSL-1 is a nonpathogenic strain of E.
coli.
Precautions of biosafety level 1:
1.
Normal laboratory personal protective equipment is generally worn, consisting of eye protection,
gloves and a lab coat or gown.
2.
Standard microbiological practices also require attention to personal hygiene, i.e., hand washing and
a prohibition on eating, drinking or smoking in the lab.
3.
Research with these agents is generally performed on standard open laboratory benches without the
use of special containment equipment.
4.
Labs are not usually isolated from the general building.
5.
Standard microbiology practices are usually enough for protection such as mechanical pipetting only
(no mouth pipetting allowed), safe sharps handling, avoidance of splashes or aerosols, and decontamination
of all work surfaces when work is complete.
6.
Decontamination of spills is done immediately, and all potentially infectious materials are
decontaminated prior to disposal, generally by autoclaving.
Biosafety level 2:
Biosafety level two deals with agents associated with human disease, in other words, pathogenic or
infectious organisms posing a moderate hazard. Example: Dealing with HIV when performing routine
diagnostic procedures or work with clinical specimens.
Precautions of biosafety level 2:
1.
Appropriate personal protective equipment (PPE) must be worn, including lab coats and gloves. Eye
protection and face shields can also be worn, as needed.
2.
Laboratory personnel have specific training in handling pathogenic agents.
3.
Access to the laboratory is limited when work is being conducted and the lab has self-closing,
lockable doors.
4.
Extreme precautions are taken with contaminated sharp items.
5.
Class II biological safety cabinet is highly recommended for work involving these agents.
Biosafety level 3:
BSL-3 is suitable for work with infectious agents which may cause serious or potentially lethal
diseases as a result of exposure by the inhalation route. Example: M. tuberculosis.
Precautions of biosafety level 3:
1.
Solid-front wraparound gowns, scrub suits or coveralls are often required.
2.
BSL-3 laboratories are located in a unique high containment building that also houses the BSL-4
laboratory that have a double-door entry.
3.
Exhaust air is not recirculated to other rooms.
4.
Standard microbiological practices are the same as for BSL-1 and BSL-2 laboratories.
5.
Class ΙΙ biological safety cabinets are suitable in BSL-3 laboratories.
6.
Additional personnel protective devices may be worn, such as respirators.
Biosafety level 4:
BSL 4 deals with extremely dangerous agents and pose a high risk of life-threatening disease.
Examples are the Ebola virus, the Lassa virus, and any agent with unknown risks of pathogenicity and
transmission.
Precautions of biosafety level 4:
1.
To the BSL 3 practices, we add requirements for complete clothing change before entry, a shower on
exit and decontamination of all materials prior to leaving the facility.
2.
Personnel must wear appropriate personal protective equipment from prior BSL levels, as well as a
full body, air-supplied, positive pressure suit.
3.
The BSL 4 laboratory contains a class III biological safety cabinet.
BSL 4 laboratories are in separate buildings or a totally isolated zone with dedicated supply and exhaust
ventilation.
Practical No 2
Isolation of genomic DNA from plants by CTAB method
Principle: extraction of genomic DNA from plant material requires cell lysis, inactivation of nucleases and
separation of desired genomic DNA from cellular debris. The cetyl trimethyl ammonium bromide (ctab)
protocol developed by muray and thompson (1980) is appropriate for extraction of genomic DNA from
plants and from plant derived food stuff and is particularly useful for the elimination of polysaccharides and
polyphenolic compounds otherwise affecting the DNA purity and quality.
Plant cells can be lysed with the ionic detergent ctab, which forms an insoluble complex with nucleic acids
in a low-salt environment. Under these conditions, polysaccharides, phenolic compounds and other
contaminants remain in the supernatant and can be washed away. The DNA complex is solubilised by
raising the salt concentration and precipitated with ethanol or isopropanol.
Extraction of genomic DNA from plants involves three steps

Lysis of cell membrane

Extraction

Precipitation
1. Lysis of cell membrane: the first step of the DNA extraction is the rupture of the cell and nucleus
wall. For this purpose, the homogenized sample is first treated with the extraction buffer containing
EDTA, tris/hcl and ctab. The lysis of the membranes is accomplished by the detergent (ctab) contained
in the extraction buffer. When the cell membrane is exposed to the ctab extraction buffer, the detergent
captures the lipids and the proteins allowing the release of the genomic DNA. In a specific salt (nacl)
concentration, the detergent forms an insoluble complex with the nucleic acids. EDTA is a chelating
component that among other metals bind magnesium. Magnesium is a cofactor for DNAse. By binding
mg with EDTA, the activity of present DNAse is decreased. Tris/hcl gives the solution a ph buffering
capacity (a low or high ph damages DNA). After the cell and organelle membranes (such as those around
the mitochondria and chloroplasts) have been broken apart, the purification of DNA is performed.
Extraction: in this step, polysaccharides, phenolic compounds, proteins and other cell lysates dissolved in
the aqueous solution are separated from the ctab nucleic acid complex. Under low salt concentration, the
contaminants of the nucleic acid complex do not precipitate and can be removed by extraction of the
aqueous solution with chloroform. The chloroform denatures the proteins and facilitates the separation of the
aqueous and organic phases. Once the nucleic acid complex has
been purified, precipitation can be accomplished.
Precipitation: in this final stage, the nucleic acid is liberated from the detergent. For this purpose, the
aqueous solution is first treated with a precipitation solution comprising of sodium acetate, which
precipitates the nucleic acid. Under these conditions, the detergent, which is more stable in alcohol than in
water, can be washed out, while the nucleic acid precipitates. The successive treatment with 70% ethanol
allows an additional purification, or wash, of the nucleic acid from the remaining salt
Protocol for genomic DNA isolation using ctab
 Take 2-3 leaves of plant tissue, homogenate it in sterilized pistil and motar with 2ml ctab buffer
(preheated at 65°c for 15 min). Ctab buffer contain (100mm tris hcl ph 8.0, 2% (w/v) ctab, 20mm
EDTA, 1.4m nacl, 1% ctab and 1% β-mercaptoethanol).
 Transfer 200 mg of homogenate to sterile eppendrof tube, and add 600 µl CTAB and mix vigorously
and incubate it in water bath at 65°c for 1hr
 Bring down the sample at room temperature, add 600µl of chloroform isoamyl alcohol mixture
(24:1), mix gently by inverting and spin at 10,000 rpm for 10 minutes
 Transfer the supernatant to a new eppendrof tube and add 600µl chroform isoamyl alcohol mixture,
mix and spin at 10,000 rpm for 10 minutes
 Collect the supernatant to new eppendrof tube and add 600µl of cold isopropanol. Mix gently to
precipitate the nucleic acid. Spin for 5-10minutes at 10,000 rpm.
 Wash the pellet with 700µl of 70% ethanol. Decant and dry the pellet at room temperature. Dissolve in 50µl
of 0.1xte+rnase. Incubate for 1 hr at 37 °c.
DNA check:
Load 2-4 µl of isolated plant genomic DNA in 1% agrose gel and determine the quality and yield.
Agarose gel electrophoresis of genomic DNA isolated from plant
Practical No 3
Extraction of genomic DNA from Animal tissue
DNA extraction from blood
The white blood cells (wbc) of peripheral blood are usually the most convenient source of human genomic
DNA for DNA analysis with respect to haemoglobinopathies. It is estimated that 10 ml of whole blood yield
approximately 250 μg of DNA, more than sufficient for complete analysis of globin genes with the methods
that are currently available (ie based on pcr). DNA is an extremely stable molecule, but enzymes which
catalyze the breakdown of nucleic acids (nucleases) are found in all cells. In intact cells the DNA is found in
the nucleus and thus is protected from the action of nucleases which are abundant in the lysosomes in the
cytoplasm. However when cells are lysed, the membranes of the cell compartments are disrupted, allowing
nucleases to come in to contact with the DNA. Thus DNA extraction uses buffers which contain inhibitors of
nuclease activity i.e.: (EDTA)
Preparation of buffers
Cell lysis buffer (clb)
The final volume will be 1000ml that contains 100mm tris-hcl (ph 7.6), 5mm mgcl2, 7mm kcl,
300mm sucrose and 1% triton x-100.
Nucleus lysis buffer (nlb)
The nucleus lysis buffer contains 100mm tris-hcl, 12mm sodium acetate/ potassium acetate or 10mm
sodium citrate, 2% sodium dodecyl sulphate (sds) and 10mm EDTA.
DNA suspending buffer (te buffer)
10mm tris-hcl and 1mm EDTA. For 1 litter
5m sodium chloride (nacl) solution
Dissolve 29.2 gm of nacl in 100ml distill water.
All the buffer must be sterilized prior use and before starting DNA extraction, chloroform and
absolute ethanol (100%) should be kept on -20oc before starting DNA extraction.
Eukaryotic DNA extraction protocol:

Take 600 μl of clb (cell lysis buffer) in pre-labeled eppendorf tube.
o Add 300 μl of EDTA/heparin blood and mix gently by inverting 4-6 times.
o Centrifuge at 4000 rpm for 5 min.
o Decant the supernatant and wash the pallet again with clb 400 μl.
o Centrifuge at 5000 rpm for 5 min.
o Repeat washing (3-4 times) until all the hemoglobin is washed out.
o Add 450 μl nlb (nuclear lysis buffer) and 100 μl of saturated nacl to dried pallet.
o Add 550 μl pre chilled chloroform. Centrifuge at 5000 rpm for 3 min. Two separate layers will
be formed.
o Withdraw the supernatant without touching the central layer to new labeled eppendorf tube. This
supernatant will be processed for further DNA extraction.
o Add 1000 μl pre-chilled ethanol.
o Place tubes at -20°c for 30 min and then centrifuge at 14000rpm for 6 min and discard ethanol
slowly.
o Add 1000 μl of 70% ethanol.
o Centrifuge at 14000rpm for 6 min. Decant ethanol and let tube dry completely.
o Add 100 μl of t.e buffer and store at 37°c for 24hrs.
o After that check DNA on 0.8% agarose gel and quantify using nanodrop.
o Store DNA at-20°c
DNA extraction buffers
Cell lysis buffer:
Tris hcl
100ml
Mgcl2
1.01g
Kcl
0.521g
Sucrose
109.54g
Triton x100 10ml
Dist. H2o
make volume up to 1000
Nuclear lysis buffer:
Tris hcl
100ml
Sodium citrate 2.9g
EDTA
20ml
Sds
20g
Dist. H2o
make volume up to 1000
T.e buffer:
Tris hcl
10ml
EDTA
2ml
Dist. H2o
make volume up to 1000
Nacl solution:
350 g in 1000ml
Tris hcl:
1. Tris
121.1g
2. Dist. H2o
make volume up to 1000
3. Hcl
make ph 7.6
EDTA solution:
1. EDTA
186.1g
Dist. H2o
make volume up to 1000
Naoh
make ph 8
What each reagent is for?
Lysis buffer:
A lysis buffer is a buffer solution used for the purpose of breaking open cells for use in molecular
biology experiments
CLB
Breaking the cell membranes open to expose the DNA along with the cytoplasm within the cell
NLB
Breaking the nucleus
Triton x-100 and sds :
Detergents Are added to break up membrane structures.
EDTA:
EDTA is responsible for chelation of divalent ions
EDTA helps to stop DNAses (DNA cutting enzymes) present in the cytoplasm from acting on the
exposed DNA
Mg2+ is an important factor for activity of DNAses. EDTA deprives the enzyme of this co-factor and
renders it inactive
DNAses are enzymes that “chew up” DNA and thus reduces the yield of genomic DNA. So it’s
important to keep them from acting on DNA of interest.
Tris-hcl:
Maintains ph
Salt solution:
The solution is treated with salt solution to make debris such as broken proteins, lipids and rna to
clump together.
Sodium chloride helps to remove proteins that are bound to the DNA.
Chloroform:
Used to help separate proteins, lipids and polysaccharides from nucleic acids in the cell. Chloroform
and water separates into two distinct phases. Lower phase will be chloroform containing
proteins,lipids and polysaccharides Chilled ethanol :
DNA doesn’t dissolve in ethanol. It will aggregate together, giving a pellet upon centrifugation
Used to separate pure DNA
Colder temperature reduces activity of enzymes that can break down DNA.
70% ethanol:
This step is to wash any residual salt away from the pelleted DNA.
T.E buffer :
Used to solubilize DNA while protecting it from degradation
Practical 4
Spectrophotometric quantification of nucleic acids
After isolation of DNA, quantification and analysis of quality are necessary to ascertain the approximate
quantity of DNA obtained and the suitability of DNA sample for further analysis. This is important for many
applications including digestion of DNA by restriction enzymes or PCR amplification of target DNA. The
most commonly used methodologies for quantifying the amount of nucleic acid in a preparation are:

Gel electrophoresis.

Spectrophotometric analysis.
Spectrophotometric analysis:
Spectrophotometer:
Instrument having two important parts:
1. Spectrometer- emit light of specific wavelength

UV (10- 400 nm)

Visible (400-780nm)
2. Photometer- measures the intensity of light
Purines and pyrmidines in nucleic acid show absorption maxima around 260nm (eg.,dATP: 259nm;
dCTP: 272nm; dTTP: 247nm). Spectrophotometers are commonly used to determine the concentration of
DNA. Inside a spectrophotometer, a sample is exposed to ultraviolet light at 260 nm, and a photo-detector
measures the light that passes through the sample. The more light absorbed by the sample, the higher the
nucleic acid concentration in the sample.
Beer-lambert law:
According to this absorbance is directly proportional to path length and concentration of solution.
A = εlc
Where:
A=absorbance in terms of optical density (od)
Ε=extinction coefficient
L=path length
C=concentration
Nucleic acid type
Value of ε
DsDNA
50g/ml
SsDNA or RNA
40 g/ml
Ss oligonucleotides
33 g/ml
Concentration of double stranded DNA in g/ml = absorbance at 260nm x 50 x dilution factor.
Concentration of single-stranded DNA and RNA in g/ml = absorbance at 260nm x 40 x dilution factor.
Sample purity:
It is common for nucleic acid samples to be contaminated with other molecules (i.e. Proteins, organic
compounds, other). The ratio of the absorbance at 260 and 280nm (a260/280) is used to assess the purity o
nucleic acids.

A ratio between 1.8 and 2.0 denotes that the absorption in the UV range is due to nucleic acids.

A ratio lower than 1.8 indicates the presence of proteins and/or other UV absorbers.

A ratio higher than 2.0 indicates that the samples may be contaminated with chloroform or phenol. In
either case (<1.8 or >2.0) it is advisable to re-precipitate the DNA.
Demerits of spectrophotometric determination:
This method is however limited by the quantity of DNA and the purity of the preparation. Accurat
analysis of the DNA preparation may be impeded by the presence of impurities in the sample or if the amount o
DNA is too little. In the estimation of total genomic DNA, for example, the presence of RNA, sheared DNA etc
Could interfere with the accurate estimation of total high molecular weight genomic DNA.
Protocol:
1. Turn the spectrometer on
2. Turn on the UV lamp 20 min before you will take your readings, the visible light lamps can be used
immediately, but UV lamp takes a while to become steady. The amount of warm up time needed
depends on the lamp and the spectrometer
3. Your sample will be DNA or RNA in water or buffer.
4. Put the sample and the blank in a matched set of quartz cuvettes
5. Set the wavelength to 260 nm
6. Blank the machine
7. Read o.d. of sample at 260 nm
8. Set wavelength to 280nm. Reblank and read the o.d. at 280
9. Calculate the concentration of DNA using following information
a. 1a260 unit of double stranded DNA= 50µg (50 µg/ml has an o.d. of 1 at 260 nm)
10. DNA concentration (µg/ml) = od260 × (dilution factor) ×50 µg DNA/ml
1od260 unit.
11. Calculate the yield of your preparation by using formula
Yield= (DNA concentration in µg/ml x total volume of solution in ml)
12. Estimate the purity of preparation by figuring the 260/280 ratios. The ratio b/w the readings at 260
nm and 280nm gives an estimate of the purity of the nucleic acid. Pure preparations of DNA should
have a 260/280 ratio of 1.8, RNA a ratio of 2.
Practical 5
Agarose Gel Electrophoresis
Principle:
Agarose gel electrophoresis separates DNA fragments according to their size. An electric current is used to
move the DNA molecules across an agarose gel, which is a polysaccharide matrix that functions as a sieve to
help "catch" the molecules as they are transported by the electric current. The phosphate molecules that
make up the backbone of DNA molecules have a high negative charge. When DNA is placed on a field with
an electric current, these negatively charged DNA molecules migrate toward the positive end of the field,
which in this case is an agarose gel immersed in a buffer bath. The agarose gel is a cross-linked matrix i.e., a
three-dimensional mesh or screen. The DNA molecules are pulled to the positive end by the current, but they
encounter resistance from this agarose mesh. The smaller molecules are able to navigate the mesh faster than
the larger ones. This is how agarose electrophoresis separates different DNA molecules according to their
size. The gel is stained with ethidium bromide so as to visualize these DNA molecules resolved into bands
along the gel. Ethidium bromide is an intercalating dye, which intercalate between the bases that are stacked
in the center of the DNA helix. One ethidium bromide molecule binds to one base. As each dye molecule
binds to the bases the helix is unwound to accommodate the stain from the dye. Closed circular DNA is
constrained and cannot withstand as much twisting strain as can linear DNA, so circular DNA cannot bind as
much dye as can linear DNA. Unknown DNA samples are typically run on the same gel with a "ladder." a
ladder is a sample of DNA where the sizes of the bands are known. Unknown fragments are compared with
the ladder fragments (size known) to determine the approximate size of the unknown DNA bands.
Approximately 10ng is visible in a single band on a horizontal agarose gel.
Materials:
•Agarose
•Tae Buffer
•Gel Casting Tray, Comb, Power Pack
•Sample DNA
•Loading Dye
•Sterile Micro Tips
•ETBR Staining Solution
•UV Transilluminator Or Gel Documentation System
Instructions:
For casting gel, agarose powder is mixed with electrophoresis buffer (TAE) to the desired concentration,
and then heated in a microwave oven until completely melted. After cooling the solution to about 60°C, it is
poured into a casting tray containing a comb and allowed to solidify at room temperature for nearly 45 min.
After the gel has solidified, the comb is removed, using care not to rip the bottom of the wells. The gel, still
in its plastic tray, is inserted horizontally into the electrophoresis chamber and just immersed with buffer
(TAE). DNA samples mixed with loading buffer are then pipetted into the sample wells, the lid and power
leads are placed on the apparatus, and a current is applied. The current flow is confirmed by observing
bubbles coming off the electrodes. DNA will migrate towards the positive electrode, which is usually
colored red. The distance DNA has migrated in the gel can be judged by visually monitoring migration of
the tracking dyes. Bromophenol blue and xylene cyanole dyes migrate through agarose gels at roughly the
same rate as double-stranded DNA fragments of 300 and 4000 bp, respectively.
Preparation of 1% agarose gel:
Weigh 1g agarose; add in 100 ml 1xTAE and melt agarose in a microwave oven for 2-3 min. Cool down to
about 45 to 50°C (bearable warmth) and pour into the gel platform with the comb in position.
Running Gel:
After solidification of the gel (approx. 45 min), place the gel in a gel tank with 1 x TAE buffer. Buffer
should be filled to the surface of the gel. Load the samples in the well and run the gel at 80V till the blue dye
runs to the end.
Staining the gel:
Prepare staining solution by adding 10 µl of 10 mg/ml stock of ethidium bromide in 100 ml of distilled
water. Place the gel in staining solution for 30 min and view the gel in UV transilluminator.
Fig 1: agarose gel electrophoresis method
Practical 6
UV Visualization and Analysis of Gel Results
If the gel is done correctly, upon visualization there will be a row of well-defined bands in the gel. However,
some errors can cause smearing, and the bands will not be distinguishable.
Gel improperly prepared
Smearing can be caused by an improperly prepared gel. If the gel is not poured correctly, it will not
polymerize or solidify evenly, thus causing the molecules to smear.
Overloading the wells
The sample of the molecule is placed into wells at one end of the gel. If these wells are filled too much, or if
the sample is not properly diluted, the excess sample may smear across the gel. In addition, if the gel is
moved after the sample is placed in the well, it can cause the sample to spill out of the well. This can also
cause smearing.
Contamination
According to "nucleic acid gel electrophoresis," another cause of smearing is contamination of the sample.
For example, a DNA sample may be contaminated with a protein, or a protein sample may be contaminated
by lipids or fats.
Determining the DNA fragment length
A 'reference ladder' can also be run in the gel. This contains a mixture of DNA fragments of known size.
Comparing the bands in your DNA sample with the bands in the reference ladder allows you to work out
how big the DNA fragments are in a particular band. DNA size is measured in base pairs (bp), or kilo-base
pairs.
Gel documentation systems
Gel documentation, or gel imaging, systems are used to record and measure labeled nucleic acid and protein
in various types of media such as agarose, acrylamide or cellulose. Systems come in a variety of
configurations depending on throughput and sample type.
Standard operating procedure (sop) for using gel documentation system (gel doc system) and handling of
agarose gels containing ethidium bromide (etbr)
1. Do not use gloves to open and close the doors of common instrument room as well as any other rooms in
bsbe.
2. Designate a separate plastic tray for carrying gel containing etbr, gloves, tissue paper and 70% ethanol /
distilled water.
3. Before using gel doc system, clean the surface of trans-illuminator with 70 % ethanol or distilled water.
4. Wear gloves to handle the gel containing etbr and then place the gel on to the surface of trans-illuminator.
5. Remove the gloves and close the door of gel doc system.
6. Document the gel picture on the computer without wearing the gloves.
7. At any moment of time, computer, keyboard, mouse and gel doc system should not be used with gloves.
8. Wear gloves to remove the gel containing etbr and clean the surface of trans-illuminator.
9. Remove the gloves and close the door of gel doc system.
10. Make an appropriate entry in the log book of gel doc system.
11. Carry all the material in the designated plastic tray.
12. Report immediately to concerned in-charge for problem regarding system, log book etc.
Practical 7
Melting kinetics of DNA
Melting temperature of DNA:
(melting temperature) temperature at which half of the DNA is melted
Melting temperature of DNA is affected by 3 main factors
 Nucleotide content of DNA molecule
 Length of DNA molecule
 Ionic strength of the DNA molecule

Refers to the separation of 2 stands
 Breaking of h-bonds + base stacking interactions




Also referred to as denaturation
Thermal energy > intermolecular bond energy (denaturation)
A:T separates more rapidly than G:C
Tm=t at which half of the DNA is melted
How to calculate melting temperature of DNA
Tm = 4(G+C) +2(A+T)
Tm =4(6) +2(5)
(suppose ATCGATATAACGGC)
Tm=24+10=34
If the GC content of a DNA molecule is 60%, what are the percentages of the four bases?
Re-naturation of DNA
Bringing the two strands together by simply lowering the temperature
 Temperature can be lower to anneal DNA strand
 Reform h-bond and base stacking
 Rapid cooling result in mismatched base pairing
 Thermal energy now too low for DNA to melt and find their proper complements
Practical No 8
Protein Estimation by Lowry method
The Lowry assay (1951) is an often-cited general use protein assay. For some time it was the method of
choice for accurate protein determination for cell fractions, chromatography fractions, enzyme preparations,
and so on. The bicinchoninic acid (BCA) assay is based on the same princple and can be done in one step,
therefore it has been suggested (Stoscheck, 1990) that the 2-step Lowry method is outdated.
Principle
The principle behind the Lowry method of determining protein concentrations lies in the reactivity of the
peptide nitrogen[s] with the copper [II] ions under alkaline conditions. Under alkaline conditions the divalent
copper ion forms a complex with peptide bonds in which it is reduced to a monovalent ion. Monovalent
copper ion and the radical groups of tyrosine, tryptophan, and cysteine react with Folin reagent to produce an
unstable product that becomes reduced to molybdenum/tungsten blue.
Equipment
In addition to standard liquid handling supplies a spectrophotometer with infrared lamp and filter is required.
Glass or polystyrene (cheap) cuvettes may be used.
Procedure:
Reagents
A. 2% Na2CO3 in 1% NaOH
B. 0.5% CuSO4 in 1 % Na-K Tartrate
Reagent-I :50ml A + 1ml B
Reagent-II :1 part Folin-Phenol [2 N]: 1 part water
BSA Standard - 1 mg/ ml
BSA is the universally accepted reference protein for total protein quantitation
Protocol:
0.2 ml of BSA working standard in 5 test tubes and make up to 1ml using
distilled water.
The test tube with 1 ml distilled water serve as blank.
Add 4.5 ml of Reagent I and incubate for 10 minutes.
After incubation add 0.5 ml of reagent II and incubate for 30 minutes .
Estimate the amount of protein present in the given sample from the
standard graph.
Measure the absorbance between 650-750 nm and plot the standard graph .
Estimate the amount of protein present in the given sample from the
standard graph.
Analysis
Prepare a standard curve of absorbance versus micrograms protein (or vice versa), and
determine amounts from the curve. Determine concentrations of original samples from the
amount protein, volume/sample, and dilution factor, if any.
Practical 9
Bradford protein assay
The Bradford assay is very fast and uses about the same amount of protein as the Lowry assay. It is fairly
accurate and samples that are out of range can be retested within minutes. The Bradford is recommended for
general use, especially for determining protein content of cell fractions and assesing protein concentrations
for gel electrophoresis.`
Principle:
The Bradford Protein Assay measures the concentration protein by adding Coomassie dye to the sample
under acidic conditions. When proteins bind with the Coomassie dye, the sample changes color from brown
to blue. The level of blue can then be measured using a spectrophotometer to determine the concentration of
protein in the sample. The assay is based on the observation that the absorbance maximum for an acidic
solution of Coomassie Brilliant Blue G-250 shifts from 465 nm to 595 nm when binding to protein occurs.
Both hydrophobic and ionic interactions stabilize the anionic form of the dye, causing a visible color change.
The assay is useful since the extinction coefficient of a dye-albumin complex solution is constant over a 10fold concentration range.
Equipment
In addition to standard liquid handling supplies a visible light spectrophotometer is needed, with maximum
transmission in the region of 595 nm, on the border of the visible spectrum (no special lamp or filter usually
needed). Glass or polystyrene (cheap) cuvettes may be used, however the color reagent stains both.
Disposable cuvettes are recommended.
Procedure
Reagents
Bradford reagent: Dissolve 100 mg Coomassie Brilliant Blue G-250 in 50 ml 95% ethanol, add 100 ml 85%
(w/v) phosphoric acid. Dilute to 1 liter when the dye has completely dissolved, and filter through Whatman
#1 paper just before use.
(Optional) 1 M NaOH (to be used if samples are not readily soluble in the color reagent).
Protocol
Warm up the spectrophotometer before use.
Dilute unknowns if necessary to obtain between 5 and 100 µg protein in at least one assay tube containing
100 µl sample
If desirred, add an equal volume of 1 M NaOH to each sample and vortex.The addition of 1 M NaOH was
suggested by Stoscheck (1990) to allow the solubilization of membrane proteins and reduce the protein-toprotein variation in color yield.Add NaOH to standards as well if this option is used.
Prepare standards containing a range of 5 to 100 micrograms protein (albumin or gamma globulin are
recommended) in 100 µl volume. See how to set up an assay for suggestions as to setting up the standards.
Add 5 ml dye reagent and incubate 5 min.
Measure the absorbance at 595 nm.
Analysis
Prepare a standard curve of absorbance versus micrograms protein and determine amounts from the curve.
Determine concentrations of original samples from the amount protein, volume/sample, and dilution factor,
if any
Practical 10
Protein separation by SDS-PAGE
Introduction:
The smaller molecules migrate faster due to less resistance during electrophoresis. The structure and the
charge of the proteins also influence the rate of migration. Sodium dodecyl sulphate and polyacrylamide eliminate the
influence of structure and charge of the proteins, and the proteins are separated based on the length of the polypeptide
chain
Principle of SDS-PAGE
The principle of SDS-PAGE states that a charged molecule migrates to the electrode with the opposite sign
when placed in an electric field. The separation of the charged molecules depends upon the relative mobility of
charged species.
Role of SDS in SDS-PAGE
SDS is a detergent present in the SDS-PAGE sample buffer. SDS along with some reducing agents function to break
the disulphide bonds of proteins disrupting the tertiary structure of proteins.
Materials Required

Power Supplies: It is used to convert the AC current to DC current.

Gels: These are either prepared in the laboratory or precast gels are purchased from the market.

Electrophoresis Chambers: The chambers that can fit the SDS-PAGE gels should be used.

Protein Samples: The protein is diluted using SDS-PAGE sample buffer and boiled for 10 minutes. A
reducing agent such as dithiothreitol or 2-mercaptoethanol is also added to reduce the disulfide linkages to
prevent any tertiary protein folding.

Running Buffer: The protein samples loaded on the gel are run in SDS-PAGE running buffer.

Staining and Destaining Buffer: The gel is stained with Coomassie Stain Solution. The gel is then destained
with the destaining solution. Protein bands are then visible under naked eyes.

Protein Ladder: A reference protein ladder is used to determine the location of the protein of interest, based
on the molecular size.
Protocol of SDS-PAGE
Preparation of the Gel

All the reagents are combined, except TEMED, for the preparation of gel.

When the gel is ready to be poured, add TEMED.

The separating gel is poured in the casting chamber.

Add butanol before polymerization to remove the unwanted air bubbles present.

The comb is inserted in the spaces between the glass plate.

The polymerized gel is known as the “gel cassette”
Sample Preparation

Boil some water in a beaker.

Add 2-mercaptoethanol to the sample buffer.

Place the buffer solution in micro-centrifuge tubes and add protein sample to it.

Take MW markers in separate tubes.

Boil the samples for less than 5 minutes to completely denature the proteins.
Electrophoresis

The gel cassette is removed from the casting stand and placed in the electrode assembly.

The electrode assembly is fixed in the clamp stand.

1x electrophoresis buffer is poured in the opening of the casting frame to fill the wells of the gel.

Pipette 30ml of the denatured sample in the well.

The tank is then covered with a lid and the unit is connected to a power supply.

The sample is allowed to run at 30mA for about 1 hour.

The bands are then seen under UV light.
Applications of SDS-PAGE
The applications of SDS-PAGE are as follows:
1. It is used to measure the molecular weight of the molecules.
2. It is used to compare the polypeptide composition of different structures.
3. It is used to estimate the purity of the proteins.
4. It is used in Western Blotting and protein ubi-quitination.
5. Analyzing the size and number of polypeptide subunits.
6. To analyze post-translational modifications.
Practical 11
Centrifugation
Centrifugation is a technique used for the separation of particles from a solution according to their size,
shape, density, viscosity of the medium and rotor speed. The particles are suspended in a liquid medium and
placed in a centrifuge tube. The tube is then placed in a rotor and spun at a define speed. Separation
through sedimentation could be done naturally with the earth gravity, nevertheless, it would take
ages. Centrifugation is making that natural process much faster. A centrifuge is a device that separates
particles from a solution through use of a rotor.
Principle
As a rotor spins in a centrifuge, a centrifugal force is applied to each particle in the sample; the particle will
then sediment at the rate that is proportional to the centrifugal force applied to it. The viscosity of the sample
solution and the physical properties of the particles also affect the sedimentation rate of each particle. At a
fixed centrifugal force and liquid viscosity, the sedimentation rate of a particle is proportional to its
size (molecular weight) and to the difference between the particle density and the density of the solution.
In a solution, particles whose density is higher than that of the solvent sink (sediment), and particles that
are lighter than it float to the top.
The greater the difference in density, the faster they move. If there is no difference in density (isopycnic
conditions), the particles stay steady.
Types of centrifuge
Low speed centrifuge
1) Most laboratories have a standard low-speed centrifuge used for routine sedimentation of heavy particles
2) The low-speed centrifuge has a maximum speed of 4000-5000rpm
3) These instruments usually operate at room temperatures with no means of temperature control. 4) Two
types of rotors are used in it,
 Fixed angle
 Swinging bucket.
5) It is used for sedimentation of red blood cells until the particles are tightly packed into a pellet and
supernatant is separated by decantation.
High speed centrifuge
1. High-speed centrifuges are used in more sophisticated biochemical applications, higher speeds and
temperature control of the rotor chamber are essential.
2. The high-speed centrifuge has a maximum speed of 15,000 – 20,000 RPM
3. The operator of this instrument can carefully control speed and temperature which is required for
sensitive biological samples.
4. Three types of rotors are available for high-speed centrifugation Fixed angle
 Swinging bucket
 Vertical rotors
Ultracentrifuge
1. It is the most sophisticated instrument.
2. Ultracentrifuge has a maximum speed of 65,000 RPM (100,000’s x g).
3. Intense heat is generated due to high speed thus the spinning chambers must be refrigerated and
kept at a high vacuum.
4. It is used for both preparative work and analytical work.
Types of centrifugation
Differential Centrifugation
The simplest form of separation by centrifugation is differential centrifugation, sometimes called differential
pelleting. Particles of different densities or sizes in a suspension will sediment at different rates, with the
larger and denser particles sedimenting faster. These sedimentation rates can be increased by using
centrifugal force. A suspension of cells subjected to a series of increasing centrifugal force cycles will yield
a series of pellets containing cells of decreasing sedimentation rate.
Practical 12
Introduction to PCR Thermocycler and Sanger Sequencing
PCR is a laboratory method used for making a very large number of copies of short sections of
DNA from a very small sample of genetic material. This process is called "amplifying" the DNA and it
enables specific genes of interest to be detected or measured.
DNA is made up of repeating sequences of four bases – adenine, thymine, guanine, and cytosine. These
sequences form two strands that are bound together in a double helix structure by hydrogen bonds (like a
spiral staircase). Each half of the helix is a complement of the other. In humans, it is the difference in the
sequence of these bases on each strand of DNA that leads to the uniqueness of each person's genetic
makeup. The arrangement of the bases in each gene is used to produce RNA, which in turn produces a
protein. There are about 25,000 genes in a human genome, and expression of these genes leads to the
production of a large number of proteins that make up our bodies. The DNA of other organisms such as
bacteria and viruses is also composed of thousands of different genes that code for their proteins.
How is the method performed?
PCR is carried out in several steps or "cycles" in an
instrument called a thermo cycler. This instrument increases and
decreases the temperature of the specimen at defined intervals
during the procedure.
1. The first step or cycle of PCR is to separate the strands of
DNA into two single strands by increasing the temperature of
the sample that contains the DNA of interest. This is called
"Denaturing" the DNA.
2. Once the strands separate, the sample is cooled slightly and forward and reverse primers are added
and allowed to bind to the single DNA strands. Primers are short sequences of bases made
specifically to recognize and bind to the section of DNA to be amplified, which are the very specific
sequence of bases that are part of the gene or genes of interest this is called Annealing. Primers
are called "forward" and "reverse" in reference to the direction that the bases within the section of
DNA are copied.
3. After the two primers attach to each strand of the DNA, a DNA enzyme (frequently Taq
polymerase) then copies the DNA sequence on each half of the helix from the forward to the
reverse primer, forming two double stranded sections of DNA, each with one original half and one
new half known as EXTENSION. Taq polymerase is an enzyme found in a bacterium (Thermues
aquaticus) that grows in very hot water, such as in geysers or hot springs. Polymerases copy DNA
(or RNA) to make new strands. The Taq polymerase is especially helpful for laboratory testing
because (unlike many other enzymes) it does not break down at very high temperatures needed to
do PCR.
4. When heat is applied again, each of the two double strands separate to make four single strands
and, when cooled, the primers and polymerase act to make four double strand sections. The four
strands becomes eight in the next cycle, eight become sixteen, and soon.
5. Within 30 to 40 cycles, as many as a billion copies of the original DNA section can be produced and
are then available to be used in numerous molecular diagnostic tests. This process has been
automated so that a billion copies of the original DNA can be produced within a few hours
Details of thermal profile for DNA amplification using PCR
General recepie and requirements
1.
2.
3.
4.
5.
6.
DNA template 50ng/ul
Primers; Forward and reverse
Taq Polymerase
dNTPs
Monovalent cations (KCL)
Divalent Cations (MgCl2)
7. PCR Buffer
8. ddH2O/PCR Water/Nuclease free water
Precautions
1. Avoid contamination
2. Tm of primers should be calculated before starting PCR
3. Conc. of template DNA should be kept in mind
Sanger Sequencing
What is sequencing?
DNA sequencing is the process of determining the sequence ofnucleotide bases (As, Ts, Cs,
and Gs) in a piece of DNA. Today, with the right equipment and materials, sequencing a
short piece of DNA is relatively straightforward.
Sequencing an entire genome (all of an organism’s DNA) remains a complex task. It requires
breaking the DNA of the genome into many smaller pieces, sequencing the pieces, and
assembling the sequences into a single long "consensus." However, thanks to new methods
that have been developed over the past two decades, genome sequencing is now much faster
and less expensive than it was during the Human Genome Project.
Sanger sequencing: The chain termination method
Regions of DNA up to about 900900900 base pairs in length are routinely sequenced using a
method called Sanger sequencing or the chain termination method. Sanger sequencing
was developed by the British biochemist Fred Sanger and his colleagues in 1977.
It’s also known as the “chain termination method,” was developed by the English biochemist
Frederick Sanger and his colleagues in 1977. This method is designed for determining the
sequence of nucleotide bases in a piece of DNA (commonly less than 1,000 bp in length).
Sanger sequencing with 99.99% base accuracy is considered the “gold standard” for
validating DNA sequences, including those already sequenced through next-generation
sequencing (NGS). Sanger sequencing was used in the Human Genome Project to determine
the sequences of relatively small fragments of human DNA (900 bp or less). These fragments
were used to assemble larger DNA fragments and, eventually, entire chromosomes.
Principle
A DNA primer is attached by hybridization to the template strand and deoxynucleosides
triphosphates (dNTPPs) are sequentially addedto the primer strand by DNA polymerase.
The primer is designed for the known sequences at 3’ end of the template strand.
M13 sequences is generally attached to 3’ end and the primer of thisM13 is made.
The reaction mixture also contains dideoxynucleoside triphosphate (ddNTPs) along with
usual dNTPs.
If during replication ddNTPs is incorporated instead of usual dNTPs in thegrowing DNA
strand then the replication stops at that nucleotide.
The ddNTPs are analogue of dNTPs.
ddNTPs lacks hydroxyl group (-OH) at c3 of ribose sugar, so it cannotmake phosphodiester
bond with nest nucleotide, thus terminates thenucleotide chain.
Respective ddNTPs of dNTPs terminates chain at their respective site. For example ddATP
terminates at A site. Similarly ddCTP, ddGTP and ddTTP terminates at C, G and T site
respectively.
Ingredients for Sanger sequencing
Sanger sequencing involves making many copies of a target DNA region. Its ingredients are
similar to those needed for DNA replication in an organism, or for polymerase chain reaction
(PCR), which copies DNA in vitro. They include:
 A DNA polymerase enzyme
 A primer, which is a short piece of single-stranded DNA that binds to the template
DNA and acts as a "starter" for the polymerase
 The four DNA nucleotides (dATP, dTTP, dCTP, dGTP)
 The template DNA to be sequenced
However, a Sanger sequencing reaction also contains a unique ingredient:
 Dideoxy, or chain-terminating, versions of all four nucleotides (ddATP, ddTTP,ddCTP,
ddGTP), each labeled with a different color of dye
Dideoxy nucleotides are similar to regular, or deoxy, nucleotides, but with one key difference: they lack a
hydroxyl group on the 3’ carbon of the sugar ring. In a regular nucleotide, the 3’ hydroxyl group acts as a
“hook," allowing a new nucleotide to be added to an existing chain.Once a dideoxy nucleotide has been
added to the chain, there is no hydroxyl available and no further nucleotides can be added. The chain ends
with the dideoxy nucleotide, which is marked with a particular color of dye depending on the base (A, T,
C or G) that it carries.
Sanger Sequencing Steps
The Sanger Sequencing method consist of six steps;
The double-stranded DNA (dsDNA) is denatured into two single-stranded DNA (ssDNA).
A primer that corresponds to one end of the sequence is attached.
Four polymerase solutions with four types of dNTPs but only onetype of ddNTP are added.
The DNA synthesis reaction initiates and the chain extends until a termination nucleotide is
randomly incorporated.
5. The resulting DNA fragments are denatured into ssDNA.
6. The denatured fragments are separated by gel electrophoresis and the sequence is determine
1.
2.
3.
4.
Procedure
1. Template preparation:
1) M13-forward-sequence copies of template strand to be sequenced must be
preparedwith short known sequences at 3’ end of the template strand.
2) A DNA primer is essential to initiate replication of template, so primer preparation
ofknown sequences at 3’end is always required.
3) For this purpose a single stranded cloning vector M13 is flanked with template
strand at 3’end which serves as binding site for primer.
2. Generation of nested set of labelled fragments:
1) Copies of each template is divided into four batches and each batch is used for
differentreplication reaction.
2) Copies of standard primer and DNA polymerase I are used in all four batches.
3) To synthesize fragments that terminates at A, ddATP is added to the reaction mixture
on batch I along with dATP, dTTP,dCTP and dGTP, standard primer and DNA polymerase
I.
4) Similarly, to generate, all fragments that terminates at C, G and T, the respective
ddNTPs i.e. ddCTP, ddGTP and ddTTP are added respectively to different reaction
mixture on different batch along with usual dNTPs.
3. GEL ANALYSIS & DETERMINATION OF DNA SEQUENCE
The last step simply involves reading the gel to determine the sequence of the input DNA.
Because DNA polymerase only synthesizes DNA in the 5’ to 3’ direction starting at a
provided primer, each terminal ddNTP will correspond to a specific nucleotide in the original
sequence (e.g., the shortest fragment must terminate at the first nucleotide from the 5’ end,
the second-shortest fragment mustterminate at the second nucleotide from the 5’ end, etc.)
Therefore, by reading the gel bands from smallest to largest, we can determine the 5’ to 3’
sequence of the original DNA strand.
In manual Sanger sequencing, the user reads all four lanes of the gel at once, moving bottom
to top, using the lane to determine the identity of the terminal ddNTP for each band. For
example, if the bottom bandis found in the column corresponding to ddGTP, then the smallest
PCRfragment terminates with ddGTP, and the first nucleotide from the 5’ end of the original
sequence has a guanine (G) base.
In automated Sanger sequencing, a computer reads each band of the capillary gel, in order,
using fluorescence to call the identity of each terminal ddNTP. In short, a laser excites the
fluorescent tags in each band, and a computer detects the resulting light emitted. Because each
of the four ddNTPs is tagged with a different fluorescent label,the light emitted can be directly
tied to the identity of the terminal ddNTP. The output is called a chromatogram, which
shows the fluorescent peak of each nucleotide along the length of the template DNA.
HOW TO READ SANGER SEQUENCING RESULTS
Reading the Sanger sequencing results properly will depend on which of the two
complementary DNA strands is of interest and what primer is available. If the two strands of
DNA are A and B and strand A is ofinterest, but the primer is better for strand B, the output
fragmentswill be identical to strand A. On the other hand, if strand A is of interest and the
primer is better for strand A, then the output will be identical to strand B. Accordingly, the
output must be converted back to strand A.
So, if the sequence of interest reads “TACG” and the primer is best for that strand, the output
will be “ATGC” and, therefore, must be converted back to “TACG”. However, if the primer
is better for the complementary strand (“ATGC”), then the output will be “TACG”, which is
the correct sequence.
In short, before starting, you need to know what you’re targeting andhow you’re going to get
there! So keeping this in mind, here is an example of the former example (TACG -> ATGC > TACG). If the dideoxynucleotides labels are T = yellow, A = pink, C = dark blue, and G =
light blue, you will end up with the short sequences primer-A, primer-AT, primer-ATG, and
primer-ATGC. Once the fragments havebeen separated by electrophoresis, the laser will read
the fragments in order of length (pink, yellow, light blue, and dark blue) and produce a
chromatogram. The computer will convert the letters, so the final sequence is the correct
TACG.
SANGER SEQUENCING VS. PCR
Sanger sequencing and PCR use similar starting materials and can be used in conjunction with
each other, but neither can replace the other.PCR is used to amplify DNA in its entirety. While
fragments of varying lengths may be produced by accident (e.g., the DNA polymerase might
fall off), the goal is to duplicate the entire DNA sequence. To that end, the “ingredients” are
the target DNA, nucleotides, DNA primer, and DNA polymerase (specifically Taq
polymerase, which can survive the high temperatures required in PCR).
In contrast, the goal of Sanger sequencing is to generate every possible length of DNA up to
the full length of the target DNA. That is why, in addition to the PCR starting materials, the
dideoxynucleotides are necessary.
Sanger sequencing and PCR can be brought together when generating the starting material for
a Sanger sequencing protocol. PCR can be used to create many copies of the DNA that is to
be sequenced.
Having more than one template to work from makes the Sanger protocol more efficient. If the
target sequence is 1,000 nucleotides long and there is only one copy of the template, it is
going to take longer to generate the 1,000 tagged fragments. However, if there are several
copies of the template, in theory it will take less time to generate all 1,000 of the tagged
fragments.
Applications
Sanger sequencing gives high-quality sequence for relatively long stretches of DNA (up to
about 900900900 base pairs). It's typicallyused to sequence individual pieces of DNA, such
as bacterial plasmids or DNA copied in PCR.
 SNP and indel genotyping.
 Find common, known and unknown sequence variations.
 Study a gene or its part.
 16 and 18S rRNA gene sequencing.
 DNA fingerprinting.
 Finds mutations and genes associated with a disease.
Practical 13
Column chromatography
What is column chromatography?
Column chromatography is described as the useful technique in which the substances
to be isolated are presented onto the highest point of a column loaded with an
adsorbent (stationary phase), go through the column at various rates that rely upon the
affinity of every substance for the adsorbent and the solvent or solvent mixture, and
are typically gathered in solution as they pass from the column at various time.
The two most common examples of stationary phases for column chromatography are
silica gel and alumina while organic solvents are regarded as the most common
mobile phases.
Column Chromatography principle
The main principle involved in column chromatography is the adsorption of the
solutes of the solution with the help of a stationary phase and afterward separates the
mixture into independent components.
At the point when the mobile phase together with the mixture that requires to be
isolated is brought in from the top of the column, the movement of the individual
components of the mixture is at various rates.
The components with lower adsorption and affinity to the stationary phase head out
quicker when contrasted with the greater adsorption and affinity with the stationary
phase. The components that move rapidly are taken out first through the components
that move slowly are eluted out last.
The adsorption of solute molecules to the column happens reversibly. The pace of the
movement of the components is communicated as:
Rf = the distance traveled by solute/ the distance traveled by the solvent
Where Rf is called retardation factor
Column Chromatography components
Components of a typical chromatographic system using a gas or liquid mobile phase
include:
Stationary phase – Generally it is a solid material having a good adsorption property
and should be suitable for the analytes to be separate. It should not cause any
hindrance in the flow of the mobile phase.
Mobile phase and delivery system – This phase is made up of solvents that
complement the stationary phase.
The mobile phase acts as a solvent, a developing agent (promotes separation of
components in the sample to form bands), and an eluting agent (to remove the
components from the column that are separated during the experiment).
Column –
For liquid chromatography: 2-50cm long and 4mm internal diameter, fabricated with
stainless steel
For gas chromatography: 1-3m long and 2-4mm internal diameter, fabricated either
with glass or stainless steel
A column’s material and its dimension are very crucial to support the stationary phase
and promote effective separations.
Injector system – Responsible for delivering test samples to the column’s top in a
reproducible pattern.
(Molecular Biology BIOT-304, BCH 305)
Detector and Chart Recorder – This gives a continuous record of the presence of the
analytes in the eluate as they come out from the column.
Detection relies on the measurement of a physical parameter (like visible or UV
adsorption).
On the chart recorder, each separated analyte is represented by a peak.
A collector at the bottom is placed at the bottom end of the column set up to collect
the separated analytes.
Column chromatography Procedure
The steps included in the column chromatography are:
Preparation of the column
Mostly the column is comprised of a glass tube with an appropriate
stationary phase
The bottom end of the column is packed with a glass wool/cotton wool or
an asbestos pad after which the stationary phase is packed.
After packing the column, a paper disc is placed on the top to avoid the
disturbance of the stationary phase during the introduction of the sample
or mobile phase.
The disturbance in the stationary phase (adsorbent layer) leads to the
irregular bands of separation.
Two types of preparing the column, known as packing techniques
namely:
Dry packing technique – The amount of absorbent needed is added as a
fine dry powder in the column and the solvent flows freely through the
column until equilibrium is achieved.
Wet packing technique – The slurry of adsorbent is prepared along with
the mobile phase and is poured into the column.
It is regarded as the ideal technique for packaging.
Page 38 of 41
(Molecular Biology BIOT-304, BCH 305)
The column should be properly washed and completely dried before inuse.
Introduction of the sample
The sample (a mixture of components) is dissolved in the minimum
amount of the mobile phase.
At one instant, the sample is introduced into the column and on the top
portion of the column, it is absorbed.
Through the elution process, the individual sample can be isolated from
this zone.
Elution technique
Through this technique, the individual components are separated
completely from the column.
The process of elution can be carried out by employing two techniques:
Isocratic elution technique – Throughout the procedure, a solvent of the
same polarity or same solvent composition is utilized.
Example: Use of chloroform alone
Gradient elution technique – Throughout the separation procedure,
solvents of gradually increased polarity or increased elution strength are
utilized.
Example: Benzene → Chloroform → Ethyl acetate → Chloroform
Detection of Components
In case the mixture separated in a column chromatography procedure are
colored compounds, then monitoring the separation progress is simple.
In case the compounds undergoing separation are colorless, then small
fractions of the eluent are sequentially collected in tubes that are labeled.
Thorugh TLC, the composition of each fraction is determined.
Column chromatography uses
Column chromatography is one of the versatile methods for
purifying and separating both solids and liquids.
Major applications:
To isolate active constituents
To separate compound mixtures
To remove impurities or carry purification process
To isolate metabolites from biological fluids
To estimate drugs in drug formulations or crude extracts
Practical #14
Thin Layer chromatography
Chromatography is an important biophysical technique that enables the separation,
identification, and purification of the components of a mixture for qualitative and
quantitative analysis.
Page 39 of 41
(Molecular Biology BIOT-304, BCH 305)
In this physical method of separation, the components to be separated are distributed
between two phases, one of which is stationary (stationary phase) while the other (the
mobile phase) moves in a definite direction. Depending upon the stationary phase and
mobile phase chosen, they can be of different types.
Thin Layer Chromatography (TLC):
Thin Layer Chromatography can be defined as a method of separation or
identification of a mixture of components into individual components by using finely
divided adsorbent solid / (liquid) spread over a plate and liquid as a mobile phase.
Components of Thin Layer Chromatography (TLC)




TLC plates
TLC chamber
Mobile Phase
A filter Paper
Principle of Thin Layer Chromatography (TLC)
TCL is based on the principle of separation through adsorption type. The separation
relies on the relative empathy of compounds towards the mobile phase and
stationary phase.The principle of TLC is the distribution of a compound between a
solid fixed phase (the thin layer) applied to a glass or plastic plate and a liquid mobile
phase (eluting solvent) that is moving over the solid phase.
Procedure of Thin Layer Chromatography (TLC)
The stationary phase is applied onto the plate uniformly and then allowed to dry and
stabilize. These days, however, ready-made plates are more commonly used.
7. With a pencil, a thin mark is made at the bottom of the plate to apply the sample
spots.
8. Then, samples solutions are applied on the spots marked on the line in equal
distances.
9. The mobile phase is poured into the TLC chamber to a leveled few centimeters
above the chamber bottom.
10. A moistened filter paper in mobile phase is placed on the inner wall of the chamber
to maintain equal humidity (and also thereby avoids edge effect).
11. Now, the plate prepared with sample spotting is placed in TLC chamber so that the
side of the plate with the sample line is facing the mobile phase. Then the chamber
is closed with a lid.
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(Molecular Biology BIOT-304, BCH 305)
12. The plate is then immersed, such that the sample spots are well above the level of
mobile phase (but not immersed in the solvent) for development.
13. Sufficient time is given for the development of spots.
14. The plates are then removed and allowed to dry.
15. The sample spots are then seen in a suitable UV light chamber, or any other
methods as recommended for the given sample.
Retention Factor (Rf ) Value
The behaviour of a compound on a TLC is usually described in terms of its relative
mobility or Rf value.
Rf or Retention factor is a unique value for each compound under the same
conditions.
The Rf for a compound is a constant from one experiment to the next only if the
chromatography conditions below are also constant:
solvent system
adsorbent
thickness of the adsorbent
amount of material spotted
temperature
.
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