Introduction:

Many tasks in molecular biology require the highly accurate transfer of small volumes of reagents. In order for the work to be meaningful, a high degree of precision is essential. Small and large volume micropipettor's are tools used to conduct transfer of liquid samples. Microfuges are small, powerful centrifuges used to separate components through centrifugation and are commonly employed for many aspects of sample processing in molecular biology.

Objective:

The objective of today's laboratory is simple: to gain competence in the highly-accurate transfer of small fluid volumes. A second objective is to gain familiarity with the microfuge as a tool for molecular biology.

Method:

Small-Volume Micropipette Technique

1. This exercise simulates the working volumes required for a small-scale reaction. Label (3) 1.5 ml tubes and add the (4) solutions to each as follows (be sure to use a separate tip to transfer each of the solutions):

Tube	Sol. I	Sol. II	Sol. III	Sol. IV
A	4 µl	5µl	1 µl	-
B	4	5	-	1
C	4	4	1	1

2. Close tops on all tubes and tap each tube on the bench top to settle the contents.
3. Place all three tubes (position balanced) within the microfuge and apply a short (several second pulse.
4. Check to see if your pipette method has resulted in an accurate 10 µl being dispensed in to each tube (set pipette to 10 µl and withdraw sample.) Is the tip filled or is sample left in the tube? Are there air bubbles? If results are not accurate repeat the exercise as required.

Large-Volume Micropipette Technique

1. This exercise simulates plasmid preparation / bacterial transformation using 100-1000µl volumes. These volumes are often far more susceptible to errors. For example, releasing the plunger too rapidly may result in air bubbles or the solution being drawn up into the piston.
2. Label (2) 1.5 ml tubes as follows and add the (4) solutions:

Tube	Sol. I	Sol. II	Sol. III	Sol. IV
E	100 µl	200 µl	150 µl	550 µl
F	150	250	350	250

1. Close tops on both tubes and tap each tube on the bench top to settle the contents.
2. Place both tubes (position balanced) within the microfuge and apply a short (several second pulse.
3. Check to see if your pipette method has resulted in an accurate 1000 µl being dispensed in to each tube (set pipette to 1000 µl and withdraw sample.) Is the tip filled or is sample left in the tube? Are there air bubbles? If results are not accurate repeat the exercise as required

Results:

1. Create a table indicating your actual measured volumes.
2. Describe (with diagrams) the method of balancing the microfuge with respect to individual sample mass as well as "position" balancing in the microfuge head.

Questions:

1. What are typical errors associated with pipette technique?
2. How would high-viscosity sample affect your micropipette technique?
3. What other tools can be used for the accurate manipulation of small-volume samples?
4. Why must the tubes be balanced within the microfuge head?

Laboratory 2

DNA Restriction Analysis and Electrophoresis

Introduction:

Restriction enzymes (endonucleases) are enzymes used to cut nucleic acids at specific target sequences. The resulting restriction fragments reveal information about the nature of the nucleic acid, often revealing the presence or absence of key sequences. The procedure involves several steps, but can be viewed as having a "restriction" enzyme treatment (to generate fragments) and agarose-electrophoresis to recover / separate fragments. Electrophoresis is followed by staining (with ethidium bromide) to visualize the fragments and the use of photography, a densitometer or a scanner to store data.

Objective:

The objective of this laboratory is simple: to treat a sample of DNA with restriction enzymes and then to separate the restriction fragments using agarose-gel electrophoresis.

Method:

Restriction of DNA

1. Add the following components to (4) labelled 1.5 ml tubes (using a fresh pipette tip for each reagent). Use this table as a summary, but follow the instructions below.

Tube	l DNA	Buffer	BamHI	EcoRI	HindIII	H2O
B	4µl	5 µl	1 µl	--	--	--
E	4	5	--	1	--	--
H	4	5	--	--	1	--
--	4	5	--	--	--	1

2. Add the 5 µl of buffer to each tube.
3. Add 4 µl of DNA to each tube.
4. Use fresh tips to add each of the restriction enzymes and distilled water.
5. Close tube tops and mix using microfuge or by tapping each tube sharply on the bench.
6. Incubate reaction tubes @ 37 ° C for at least 20 minutes.

Cast 0.8% Agarose Gel

1. Seal ends of the casting tray with tape and insert the well-forming "comb".
2. Carefully pour enough agarose to fill tray to about 5 mm depth. (Gel should cover ~ 1/3 the height of the comb teeth. Use a pipette tip to remove large bubbles while the gel is still liquid. Gel will become cloudy and solidify in about 10 minutes.
3. When gel has set, unseal ends of casting tray and place tray on platform of gel box such that the comb is at the negative (black) electrode.
4. Fill the box with TBE buffer to just cover the surface of the gel.
5. Gently remove the comb (by pulling straight up), being extremely careful not to rip the wells.
6. Make sure the wells are just under the surface of the buffer.

Agarose gel Electrophoresis

1. Add loading dye to each reaction tube (1 µl dye to each tube, mix by rapping on bench-top).
2. Use a micropipettor to load the contents of each reaction tube into a separate well on the gel.Mix or briefly spin with microfuge to ensure all liquid is at the bottom of the microtubes. Use a fresh tip for each reaction mixture. Make a table recording the location of each sample.
3. After loading all samples, close the electrophoresis box and hook up the leads to the power supply. Set supply to 100-150 Volts. The ammeter should read 50-100 milliamperes.
4. Electrophorese for 40-60 minutes. (Good separation is indicated by the bromophenol blue band moving 4-8 cm from the wells. Stop the electrophoresis before the blue bands runs off the gel.
5. Turn off the power supply, disconnect the leads and remove the top of the electrophoresis box.

Gel Staining (Ethidium Bromide Method)

1. Flood gel with ethidium bromide (1µg/ml) and allow to stain for 5-10 minutes.
2. After staining, decant off as much stain as possible.
3. Rinse gel (and tray) under running tap water.
4. View gel under ultraviolet light source to see bands.

Gel Staining (Methylene Blue Method)

5. Flood gel with 0.025% methylene blue and allow to stain for 20-30 minutes.
6. After staining, decant off as much stain as possible.
7. Rinse gel (and tray) under running tap water. Allow to soak in water to remove excess stain.
8. View gel over light box or capture image with a scanner.

Results:

1. Make an accurate schematic diagram showing the locations of the restriction fragments on your gel.. The base pairs observed for the Hind III fragments should include bp. Of : 27,491; 23,130; 9,416; 6,557; 4,361; 2,322; 2,027; 564; 125.
2. Based on the measurements of your gel, identify the fragment sizes for your restriction products.
3. Make a table listing the fragments recovered from each restriction enzyme.

Questions:

1. Why is there sucrose in the loading dye?
2. What would occur if the box were filled with water rather than buffer?
3. What would occur if the electrodes were reversed?
4. How is this technique useful in identifying individual humans?
5. How many restriction fragments are generated from a eight-site circular and linear DNA molecule?

Laboratory 3

Modification of DNA through Methylation

Introduction:

In E.Coli, the restriction enzyme EcoRI recognizes a specific sequence. Alternatively, a second enzyme, EcoRI methylase serves to methylate (add a methyl group) to two adenines in the target sequence which serves to protect the DNA from restriction: in essence this is a protection of the cells own DNA from the EcoRI digestion. Methylation is therefore one of the tools the molecular biologist can employ to direct restriction analysis away from specific sequences.

Objective:

The object of this laboratory is to conduct methylation modification of l DNA with EcoRI methylase and perform a restriction digestion of the modified DNA. The results will be examined through agarose gel electrophoresis.

Method:

Methylation of DNA

1. Add the following components to (4) labelled 1.5 ml microtubes (using a fresh pipette tip for each reagent). Use this table as a summary, but follow the instructions below.

Tube	l DNA	Buffer/ SAM*	Water	EcoRI methylase
1	4µl	5 µl	2 µl	--
2	4	5	1	1µl
3	4	5	1	--
4	4	5	--	1
5	4	5	1	--
6	4	5	--	1

*Buffer/SAM (200µl restriction buffer (10X) 10µl of 30mM

S-adenosyl methionine in 4.5mM sulfuric acid with

790µl of deionized water.

1. Add components, but use fresh tips to add each of the different reagents.

1. Close tube tops and mix using microfuge or by tapping each tube sharply on the bench.
2. Incubate reaction tubes @ 37 ° C for at least 20 minutes.

Cast 0.8% Agarose Gel

1. Seal ends of the casting tray with tape and insert the well-forming "comb".
2. Carefully pour enough agarose to fill tray to about 5 mm depth. (Gel should cover ~ 1/3 the height of the comb teeth. Use a pipette tip to remove large bubbles while the gel is still liquid. Gel will become cloudy and solidify in about 10 minutes.
3. When gel has set, unseal ends of casting tray and place tray on platform of gel box such that the comb is at the negative (black) electrode.
4. Fill the box with TBE buffer to just cover the surface of the gel.
5. Gently remove the comb, being extremely careful not to rip the wells.
6. Make sure the wells are just under the surface of the buffer.

Restriction Enzyme Reaction

1. Add the following components to (4) labelled 1.5 ml microtubes (using a fresh pipette tip for each reagent).

Tube	EcoRI	HindIII
1	-	-
2	-	-
3	1 µl	-
4	1 µl	-
5	-	1 µl
6	-	1 µl

2. Close tube tops and mix using microfuge or by tapping each tube sharply on the bench.
3. Incubate reaction tubes @ 37 ° C for at least 20 minutes.

Agarose gel Electrophoresis

1. Add loading dye to each reaction tube (1 µl dye to each tube, mix by rapping on benchtop).
2. Use a micropipettor to load the contents of each reaction tube into a separate well on the gel. Use a fresh tip for each reaction mixture. Make a table recording the location of each sample.
3. After loading all samples, close the electrophoresis box and hook up the leads to the power supply. Set supply to 100-150 Volts. The ammeter should read 50-100 milliamperes.
4. Electrophorese for 40-60 minutes. (Good separation is indicated by the bromophenol blue band moving 4-8 cm from the wells. Stop the electrophoresis before the blue bands runs off the gel.
5. Turn off the power supply, diconnect the leads and remove the top of the electrophoresis box.

Gel Staining (Ethidium Bromide Method)

1. Flood gel with ethidium bromide (1µg/ml) and allow to stain for 5-10 minutes.
2. After staining, decant off as much stain as possible.
3. Rinse gel (and tray) under running tap water.
4. View gel under ultraviolet light source to see bands.

Gel Staining (Methylene Blue Method)

1. Flood gel with 0.025% methylene blue and allow to stain for 20-30 minutes.
2. After staining, decant off as much stain as possible.
3. Rinse gel (and tray) under running tap water. Allow to soak in water to remove excess stain.
4. View gel over light box or capture image with a scanner.

Results:

1. Create an accurate schematic diagram (or captured image) of your electrophoresis results.
2. Discuss the effects of methylation on the results. How could this method be used advantageously?

Questions:

1. What is the purpose of samples 1, and 2? Why are these samples necessary?
2. What is the advantage of a host cell being able to modify any restriction recognition sites on its' own DNA?
3. What adaptive value do methylases have for bacteria?
4. How are methylases used in constructing a genomic library?
5. What is the role of DNA methylation in higher organisms?

Laboratory 4

Transformation of E.coli with Plasmid DNA

Introduction:

Transformation refers to the insertion of a specific genetic sequence into the genome of a host DNA. There are essentially four separate stages: pre-incubation (@ 0° C to freeze cell membranes and distribute charged phosphates), incubation (to add DNA @ 0° C), heat-shock (a brief incubation @ 42° C to allow plasmids to enter the host cells) and recovery (incubation @ 37° C in an appropriate broth). There are many variations on this type of procedure and others continue to be developed,; but a basic understanding of the principles is essential for the molecular biologist.

Objective:

To conduct a "rapid" transformation of E.coli cells using plasmid DNA to gain an understanding of the aseptic methodology and many other considerations required in transformation procedures.

Method:

Note: aseptic technique is critical for all aspects of the following procedures.

1. Aseptically transfer 250 µl of CaCl₂ solution (50 mM) to (2) sterile 15 ml tubes. One will be the control ( - pAMP) sample, the other will be the treated (+ pAMP) sample. Place both samples on ice.
2. Use a sterile loop to transfer one or two large E.coli colonies to the + pAMP tube. Swirl the loop / needle to dislodge the cells into the CaCl₂ solution. Suspend the cells in the solution by aseptically micropipetting the suspension several times into an appropriate micropipette. Close the tube and examine to ensure even distribution of the cells. Place the suspension on ice.
3. Repeat step (2.) for the - pAMP solution.
4. Use a micropipette to add 10 µl of a 0.005 µg/µl pAMP solution directly into the + pAMP tube. Tap to mix, but avoid making bubbles or getting solution up the tube walls. Place the sample back on ice and leave both sample on ice for an additional 15 minutes. While waiting label two LB plates (one + control, one - control) and two LB/amp plates ( one + and one -)
5. Heat shock the cells by removing tubes from ice and immediately immersing in 42° C water bath for 90 seconds. Immediately after the heat shock return cells/tubes to ice for at least one minute.
6. Place both tubes in a rack at room temperature.
7. Aseptically transfer 250 µl of LB broth to each tube. Tab to mix.
8. Spread each type of cell suspension on the plates using the following table as a guide:

	Transformed cells + pAMP	Non-transformed - pAMP
LB / amp	100 µl	100µl
LB	100 µl	100 µl

(Use appropriate sterile microbiological technique to spread the samples on the appropriate plate surface.)

9. Incubate the inoculated plates face-down @ 37° C for 24 hours.
10. The next day count your colonies on each type of plate.
11. Inoculate appropriate media (LB broth) to maintain a culture of both the control and transformed populations for next weeks lab.

Results:

1. Prepare a flow chart of the transformation procedure.
2. Determine the transformation efficiency (expressed as the number of antibiotic-resistant colonies per µg of pAMP DNA. (The idea is to determine the mass of pAMP which was spread on the experimental plate and was therefore responsible for the resulting transformations.

To calculate transformation efficiency:

	determine the total mass of pAMP (conc. X vol. = mass).
	determine the volume of cell suspension loaded onto the + LB/amp plate (vol. Susp. Spread / total vol. Suspension).
	determine the mass of pAMP in cell suspension spread onto plate: total mass pAMP x fraction spread = mass pAMP spread.
	Finally determine the number of colonies / µg of pAMP. Express as colonies observed / mass of pAMP spread = transformation efficiency.

Questions:

1. What factors affect transformation efficiency?
2. Why do you have positive and negative controls in both the plate/media types?
3. What other transformation methodologies are available?

 Laboratories 5 - 6

Isolation and Identification of Plasmid DNA

Introduction:

During the last laboratory transformed E.coli were identified through resistance to ampicillin. This confirms the presence of the plasmid DNA but offers no insight as to the structure of the plasmid. A standard method of characterizing the plasmid DNA is to isolate it and conduct a restriction analysis / electrophoresis to determine the nature of the plasmid DNA. This involves the growth of the culture (to provide sufficient DNA for analysis) followed by cell lysis (to release cellular contents), separation to remove non-DNA components and restriction analysis / electrophoresis to visualize the nature of the isolated plasmid.

Objective:

This two-laboratory exercise will involve the isolation of the plasmid DNA (today's exercise) and the subsequent restriction analysis and electrophoresis of the isolate (next laboratory period).

Method:

1. Label (2) microtubes (1.5 ml) as "R" (resistant) and "S" (sensitive). Transfer 1 ml of the overnight cultures into the appropriate tube. (i.e. amplicillin resistant in "R" tube).
2. Ensure the tubes balance and centrifuge for one minute to recover the cells. Remove as much supernatant as possible without disturbing the cells.
3. Add 100 µl of glucose / TRIS / EDTA solution (GTE) to both tubes. Agitate appropriately to suspend the cells in the GTE solution.
4. Lyse the cells in each tube by adding 200 µl of SDS-NaOH solution and mix each tube.
5. Place the suspensions on ice for (5) minutes (as the cells lyse the solution should go from turbid to relatively clear).
6. Precipitate the proteins by adding 150 µl of ice-cold potassium acetate / acetic acid (KOac).
7. Place on ice for (5) minutes then centrifuge for 5 minutes to remove proteins.
8. Transfer 400 µl (of the supernatant only) from each tube, making sure no precipitate is transferred into new (labeled) tubes. Discard the precipitate.
9. Precipitate the DNA by adding 400µl of isopropanol. Mix each tube and allow two minutes at room temperature for precipitation.
10. Centrifuge for (5) minutes to recover a dense DNA pellet. Remove as much supernatant as possible but leave the DNA intact. Wash the DNA with 200 µl of ethanol (mix to re-suspend DNA and then centrifuge (2-3) minutes to recover DNA).
11. Remove the supernatants and keep the DNA pellets. Dry the pellets in open air (i.e. leave tubes open to evaporate ethanol). All ethanol must be removed before proceeding.
12. Add 15 µl of TE (TRIS / EDTA) solution to both tubes.
13. Freeze DNA solutions until ready for restriction and electrophoresis.

 

Restriction Analysis and Electrophoresis

1. Set up four minitubes as follows, adding the restriction enzyme mixture (Bam HI / Hind III) as a last step.

	pAMP	S1	R1	Control
pAMP DNA	5 µl	-	-	5 µl
S1 DNA	-	5	-	-
R1 DNA	-	-	5	-
Buffer/Rnase	5	5	5	5
Water	5	5	5	5
Bam/Hind Mix	5	5	5	-

2. Perform the restriction digestion, electrophoresis and staining / photography as in laboratory (2), etc.

Results:

1. Make an accurate schematic diagram showing the locations of the restriction fragments on your electrophoretic gel..
2. Based on the measurements of your gel, identify the fragment sizes for your restriction products.
3. Make a table listing the fragments recovered from each restriction enzyme.
4. Determine the approximate mass of DNA you have recovered.

Questions:

1. What is the purpose of the GTE buffer?
2. Which steps of the isolation remove proteins / lipids / DNA?
3. What would you expect to observe in your transformed cells?
4. Does your data indicate the presence of pAMP in transformed cells? Why?
5. Does your data indicate the presence of pAMP in non-transformed cells? Why?

Laboratories 7 -8

PCR - Polymerase Chain Reaction

Introduction:

Since its introduction in 1985, the polymerase chain reaction (PCR) has become an extremely useful tool for molecular geneticists / biologists. PCR allow scientists to replicate and amplify a specific nucleic aid sequences up to a 1,000,000 times in only a few hours. The reaction is simple and fast and has many applications. The key component is the enzyme DNA polymerase which is essential for the amplification. PCR employs several key steps: heating to 94-96° C to denature (separate the two-strands of the DNA helix), annealing at 50-65° C (to anneal / hybridize primers to complementary sequences) and several minutes @72° C (for the polymerase to ad complementary DNA to the primers). After thirty such cycles there can be as many as a billion exact copies of the original sequence produced.

The key development for the procedure was the discovery of a heat-stable polymerase: able to function after the numerous heating cycles. This critical enzyme was recovered from a bacteria found at hot springs: Thermus aquaticus.

Objective:

In this two-laboratory exercise a sequence of l DNA will be amplified using an automated thermal cycler, while in the second laboratory the amplified DNA will be examined through agarose gel electrophoresis.

Method:

1. The automated thermal cycler must be set to: 96° C, 1 minute, 58° C 1 minute, 1 cycle – link to - 96° C 30 seconds, 58° C 1 minute, 16 cycles – link to - 58° C 10 minutes.
2. Label a 1.5 ml microtube as "master reaction". Add 45m l of l DNA, 90 m l of PCR mixture and 90 m l of MgCl₂.)
3. Label (4) 0.5 ml microtubes as:

0 cycles (control)

9 cycles for Et. Br. Or 18 for methylene blue

13 cycles for Et. Br. Or 22 for methylene blue

17 cycles for Et. Br. Or 26 for methylene blue

4. Add 50 m l of reactants from the master reaction tube to each of the labeled PCR tubes.
5. Add one drop of mineral oil to each of the reactant tubes (be careful not to touch dropper to PCR reactants or tube).
6. Place the PCR tubes on ice. The control (0 cycles tube) will remain on ice during the thermal cycling procedure.
7. Programming of the automated thermal cycler will be done by the instructor 9as described earlier).
8. Load the reaction tubes into the thermal cycler and start the program.
9. After 4 cycles have been completed place the 13-cycle tube(s) into the thermal cycler (do not touch the heating block).
10. After 8 cycles have been completed place the b9-cycle tubes into the heating block (do not touch the heating block).
11. When cycles are complete, recover your samples and maintain on ice (refrigerated) until the electrophoretic analysis.
12. Prepare a 1.0 % agarose gel as previously described
13. Label (4) 1.5 ml microtubes as: 0 cycles (control), 9 cycles, 13 cycles and 17 cycles.
14. Transfer 20 m l of loading dye to each 1.5 tube. Close top and tap or micropipette to mix.
15. Add 20 m l of each PCR/loading dye sample into adjacent wells of the 1 % agarose gel.
16. Add (2) m l of loading dye to a 20 m l aliquot of DNA size markers and load one 20 m l sample of this solution per gel.
17. Electrophorese at 100 volts for around 40 minutes. Adequate separation will have occurred when the bromophenol blue dye front has moved 40-50 mm from the wells.
18. Turn off the power supply, remove the casting tray and transfer gel to an appropriate staining tray.

Results:

1. Make an accurate reproduction of the stained gel. Report if the amplified samples show greater intensity than the other samples.
2. Attempt to determine the size the the PCR product in relation to the DNA size markers. A good method is to plot migration as a function of molecular weight:

where D (distance) = 1 / log MW

3. Plot distance migrated / base pair length for each marker fragment.
4. Use the resulting line to infer PCR sample MW.

Questions:

1. Use the following information to determine how many DNA molecules were present in 2.5 ng of l DNA added to each PCR reaction: { one molecuyle of l DNA has 48,502 b.p., one b.p. has an average atomic mass of 650 g / mole, there are 6.02 x 10²³ molecules / mole.}
2. How many copies of the PCR product are produced after 17 amplification cycles?
3. What is the threshold of detection with the stain you employed?
4. Describe a minimum of (5) applications of PCR methodologies.

01/04/2010