00 Pre-Laboratory Notes

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Cell & Molecular Biology Laboratory: MKCCanlas Page 1 of 13Biology Department, School of Science & Engineering

Ateneo de Manila University

General Instruction & Laboratory Methods

Lab Rules

1. Always be prepared before coming to the lab. Read and understand the laboratory manual before

performing the exercises.

2. Bring to the lab bench only your lab notebooks, manual and other necessary items for the experiment.

Leave bags and personal belongings at assigned places only.

3. Wear appropriate clothing and protective gear appropriate to the situation: safety glasses, lab gown,

gloves, mask, etc. If you are wearing gloves, always remember that pens, doorknobs, or anything you

touch may potentially be contaminated with whatever your gloves have come into contact with.

4. Work in designated areas. Dispose of materials as indicated by your instructor

5. Use sharp and breakable objects properly. Carry sharp objects (syringe with needle, Pasteur pipettes,

etc.) with the sharp ended pointed down and away from you. Do not leave pipette tips protruding

from the lab bench into the aisle.

6. Wash your hands before starting an experiment and before leaving the lab.

7. Eating, drinking, smoking, applying make-up is strictly forbidden in the lab.

8. Observe good laboratory practice and personal hygiene. Keep your work area clean and uncluttered.

9. Report dangerous activities and situations. Pay attention to the way other people are conducting

themselves in the lab. Someone elses mistakes or carelessness can hurt you. Be aware of yoursurroundings at all times.

10. If you dont know or are not sure ASK your instructor or any member of the lab staff.

Chemical & Physical Safety

1. Handle chemicals and equipment with care. Be aware of locations of showers, eyewashes, fume

hoods and first aid assistance.

2. Know what you are handling, several chemicals used in the laboratory are hazardous. Refer to Material

Safety Data Sheets (MSDS) supplied by manufacturers or on the Web.

Take note especially of the following:

Phenolcan cause severe burns

Acrylamidepotential neurotoxinEthidium Bromidemutagen

These and other chemicals are not harmful is used properly: always wear gloves when using potentially

hazardous chemicals and never mouth pipette them.

3. If a spill occurs, immediately call the lab instructor or lab staff member. Remove contaminated clothing

immediately. Flush skin with water for at least 5 minutes, wash with mild soap and water. Seek medical

advice. Contain spilled material in appropriate containers and evaluate the area if necessary.

4. Dispose of all reagents properly. Halogenated, organic, heavy metal, ethidium bromide and solid

waste will have proper disposal containers.

5. Never operate equipment you are not familiar with without first consulting the users manual. If the

manual is not available, ask the lab instructor or a member of the lab staff for assistance. Be extra

careful when plugging in equipment. Always check the voltage requirement; take not is 110- or 220 V.

Use a transformer when necessary.

Voltage used for electrophoresis is sufficient to cause electrocution. Cover buffer reservoirs duringelectrophoresis and always turn of the power supply and unplug the power source before removing the

gel.

6. General Housekeeping. All common areas should be kept free of clutter and all dirty glassware should

be dealt with appropriately. Keep your work areas clean. Since you will use common facilities, all

solutions and everything stored in an incubator, refrigerator, etc., must be labeled. In order to limit

confusion, each person should use his initials or another unique designation for labeling plates, etc.


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Unlabeled material found in the refrigerators, incubators, or freezers may be discarded. Always mark the

backs of the plates with your initials, the date, and relevant experimental data.

Biosafety

1. Always handle microbial and mammalian cell cultures as if they were infectious agents or pathogens.

Always wash your hands before leaving the laboratory.

2. Never discard cultures in the sink. All cultures are counted as biohazards and must be decontaminated

by autoclaving.

3. Place materials and cultures for decontamination in biohazard bags prior to autoclaving.

4. If a bacterial or cell sample spill occurs, immediately pour disinfectant (5% Lysol solution) over the spill.

Wipe with paper towels and dispose paper towels as biohazard waste.


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Laboratory Notebooks and Laboratory Reports

Lab Notebook. An observation notebook should be kept for laboratory experiments.

Always bring lab notebook. A lab notebook is a 100-page hard-bound record book with page numbers.

Ring bound and loose leaves are not allowed. Use blue or blank ink. Number each page at the top righ

corner. Do not use pencils for writing data on the notebook. Mistakes should not be erased; they should

be crossed out with a single line. The notebook should always be up-to-date and may be collected by the

instructor at any time.

Write your name on the inside front cover of your lab notebook. The first page (page 1) will be designated

for the table of contents which will be continuously updated throughout the semester in the following

format:

Date/s Performed Experiment Page No.

Start your data entry on page 5. For each exercise (start new experiments on new pages), an entry is

expected in your lab notebook with the following format:

1. Experiment Title:2. Date/s performed:3. Outline/Flowchart of Procedure:4. Reagent Preparations (if necessary):5. Other Calculations (if necessary):6. Notes/Reminders (if any):7. Results:

Items 13 are expected to be filled out before class hours. Lab notebooks will be subject to spot-checks

during lab period.

Write everything that you do in the laboratory in your lab notebook. The notebook should be organized byexperiment and should not be organized as a daily log.

Buffer compositions (unless standard or included in a kit), biological reagents and special equipment are

noted.

All results for the experiment are expected to be found in your notebook. These may be in the form o

observations, data tables, drawings or photographs (pasted). All should be PROPERLY labeled with the

date, identification and source.

Lab Reports. A well-researched paper summarizing the each experiment and the results will be submitted.

Formal lab reports (group report, unless otherwise stated) must be submitted a week after the exercise isfinished. Late lab reports will not be accepted. Automatic zero points for that exercise. Deadlines shall be

strictly followed. Points will be deducted for not following directions. Plagiarism is a major offense and

subject to sanction or disciplinary action.

If a student is absent during the time the experiment is performed, he/she may contribute to the lab report,

however, 25% will be deducted from the lab report grade for his/her individual grade.


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The formal report should be computerized an printed on short bond paper, single spaced, font Arial 11, 1

inch margin on all sides. The format will be as follows:

Group # Date Submitted:Group members: Date Performed:

Experiment No.

Experiment Title

Abstract

The abstract is a condensed version of the entire report summarizing the purpose and objectives of the experiment,a short description of the methodology, results and major conclusions; all stated in a single paragraph and in the pastense.

I. Introduction

Very brief background of the exercise, its nature and scope

Information pertinent to the execution of the experiment or interpretation of results is stated.II. Methodology

Methods should not be copied exactly from the lab exercise but stated briefly in paragraph form and in anorganized manner

Usually presented in chronological order and stated in the past tense Tables may be usedif the experiment includes a set of reactions with multiple variables.

III. Results & Discussion

Most important part of the report Description and explanation of significance of the materials and methods used

Incorporate data into the text (presented as tables, graphs, figures, photos) as you discuss. Figuresshould be captioned and lebelled properly!

Analysis and interpretation of data and results

Compare data with expected results or theoretical information. Account for differences orinconsistencies.

Answers to guide questions may be incorporated here.IV. Conclusions & Recommendations

Summarize results

Draw conclusions based on the results and make applicable recommendationsV. References

State reference materials that were actually used

References will be presented alphabetically regardless of source type (book, journal, internet, abstract,unpublished, etc.

Include at least 3 book or journal references. Have minimal internet-based sources.


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Using Micropipettors

Molecular biology laboratories make frequent use of a volumetric measuring device called a micropipettor,which is made to deliver precise, accurate volumes in the range of 0.2 to 1000 microliters (L)in the hands of a

skilled operator. Hence, it is important to learn how to operate this instrument properly because otherwise it wil

not deliver the correct volume. Moreover, each device costs about PhP 20,000.00 or more, and if misused

costly repairs or replacement will be required. So take some time to learn how to do it right!

Metric Conversions

Familiarize yourself with metric units of measurements and their conversions. This exercise uses volume

measurements (base unit: liter) but the prefixes will also apply to mass (base unit: gram) or linear measurements

(base unit: meter).

1 mL = 0.001 liter or 1/1,000 liter 1,000 mL = 1 liter1 L = 0.000001 liter or 1/1,000,000 liter 1,000,000 L = 1 liter

1 L = 0.001 mL or 1/1,000mL 1,000 L = 1 ml

CAUTION: No-nos when using micropipettors

1. Do NOT use without putting a tip on the pipettor; the barrel should never touch the liquid, only the tip.

2. Do NOT lay down the pipettor when its tip contains liquid. Keep it vertical!

3. Do NOT attempt to set the volume above or below the stated range of the device. Pay careful

attention to the volume scale and the range!

4. Do NOT let the plunger snap back after taking up or ejecting liquid this damages the piston. Maintain

steady control of the plunger speed.

Familiarizing yourself with micropipettors

1. Examine your set of micropipettors. We will be using as many as five different pipettors and they shouldcover the range from 0.2 to 1000 microliters (L). Sometimes the pipettors (and also the accompanying

tips) are color-coded by size, clear or white for smallest, yellow for mid-range, blue for largest volume,

but this is not always the case. Look at your set for the presence or absence of color-coding on each

pipettor.

2. Locate working parts. Pick up any one micropipettor. Identify (but do not manipulate yet) the plunger

on the top of the device, the digital volume readout on the body, the separate tip ejector plunger, and

the shaft that accepts the tip.

3. Insert a tip onto the shaft of the pipettor.

a. Get the correct tip to fit your pipettor. Color coding may help but is not always present (e.g. yellow

tip to pipettor with yellow top).

b. Press firmly to create an airtight seal between the barrel of the shaft and the tip so the instrument

can do its work in drawing up the precise volume of liquid.

Never insert a pipettor into liquid without a tipthis would ruin the piston that measures the precise

volume.

4. Practice operating the plunger on the top of the device as follows: (we are not involving liquid yet)

a. Place your thumb on the plunger. Use the rest of your fingers to grip the instrument with your index

finger resting in the slot.

b. Gently, with steady pressure, depress the plunger till you reach the first stop, that is, till you can

detect resistance. Then depress further, till you get to the second stop. (One version of a


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micropipettor might have a third stop which serves to eject the tip, but most versions have only

the two stops with a separate tip ejector.)

c. Repeat a few times to get the feel of the stops and to practice steady control of plunger speed.Never let the plunger snap back because this can damage the piston.

5. Setting the Volume. These pipettors are termed variable volume because you can set the volume to

a value within the range of the instrument. Each pipettor has its particular range of values. Examine

your pipettor set and know the volume range of each pipettor:

pipettorVolume range (L)

tipminimum maximum

P2 0.2 2.0 colorless

P10 0.5 10.0 colorless

P20 2.0 20.0 yellow

P100 20.0 100.0 yellow

P1000 200.0 1000.0 blue

Locate the volume adjustor: sometimes this is the plunger itself or sometimes it is a knob near the top of

the pipettor.

Next, look at the volume readout window; different brands of pipettors have different approaches tothe readout scale. Your pipettor may have a direct digital readout with the decimal place clearly

indicated for ease of use or it may have a micrometer readout setting that varies with the size of the

pipettor. A direct readout is easy, no need to explain, the decimal points are shown. But the

micrometer readouts require more attention. See appendix A for detailed directions if you are using

one of these pipettor sets; tricky!

Do not attempt to set the volume beyond the pipettes minimum or maximum value doing so

damages the gears. That is why it is so important to know how to read the range and volume readout

of each kind of pipettor.

P100

0

P100

P20

P10

P2

1 1 1

7 0 5 8 7

5 0 5 5 5

750L

100L

15.5L

8.5L

1.75L

Digital display for Gilson Pipetman micropipettors

DO NOT DIAL PAST THE LIMITS OF THE PIPETTE!

Using micropipettors: Working with Sample Liquid.

1. First arrange your materials on your workspace for easy access, and sit at a height so that you can rest

your elbow on the work surface.

2. Pick up a micropipettor with a tip in place. If it is the small-volume pipettor, set the volume to 4 L. If it is

the mid-range pipettor, set the volume to 75 L. If the largest volume pipettor, set the volume to 350 L.

3. Obtain a microcentrifuge tube containing colored liquid sample. Hold the opened tube firmly in one

hand at nearly eye level to observe the movement of the liquid as you operate the pipettor.


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4. Hold the micropipettor in a vertical position over the tube.

5. To withdraw sample from the tube:

- depress the plunger to the first stop and hold. (This action displaces air equal to set volume from

the tiphence, the term an air-displacement pipettor.)

- immerse the tip of the pipettor into the sample liquid. Do not push the tip into the very bottom of

the tube as this may restrict withdrawal of the sample liquid.

- Slowly release the plunger to draw up the liquid into the tip.

- Pause for a second or two to allow full intake of liquid into tip.

- Remove tip from sample liquid. (Some recommend sliding the tip along the inside of the tube,

others say to pull the tip straight out).

- Examine the liquid in the tip to be sure that there is no air space at the end of the tip. (There will

be, of course, an air space on top where the tip meets the pipettor shaft. This cushion of air

protects the pipettor itself from being contaminated by the sample.)

- If there is an air space on bottom, eject liquid sample and repeat sample uptake steps till there is

no visible air space at the bottom of the tip .

- To assure maximum accuracy, you can examine the outside of the tip and wipe off liquid

droplets that cling to outside, taking care to stay away from tip outlet to avoid removing liquid

from the tip interior.Always keep the pipettor in an upright position while there is liquid in the tip, so that the liquid does

not contaminate and damage the shaft of the pipettor. 6. To deliver sample into a new tube:

- Insert the tip of the pipettor into the inside wall of the new tube. This creates a capillary effect to

help remove sample completely from the tip.

- Slowly depress the plunger to the first stop, then on to the second stop to release the last bit of

sample liquid. Pause for a second with plunger in depressed position to allow time for full

release.

- Keep the plunger depressed while you remove the tip from the new tube out of the liquid ( this

avoids withdrawing liquid back into the tip!)

- Next,Release plunger gradually to resting position.If you touch the tip into a liquid mixture, be sure to use a fresh tip for the next sample delivery to

avoid cross-contamination of reagents.

7. Eject tip into discard container for later disposal.

8. Practice this technique with all pipettors in your set to get the feel of each pipettor and get a visual

picture of volumes in the pipettor tips.

Practice!Take the 0.5-10 L micropipettor, put on appropriate tip and

1. Take a piece of parafilm* and place 5 individual 10 L drops of water on it.

2. Transfer 5 L of colored fluid from stock into the first drop. Mix.

3. Transfer 4 L of colored fluid from the first drop into the second drop. Mix

4. Transfer 3 L of colored fluid from the second drop into the third drop. Mix.

5. Transfer 2 L of colored fluid from the third drop into the fourth drop. Mix.6. Transfer 1 L of colored fluid from the fourth drop the fifth drop. Mix.

10 l is approximately the size of __________________.

Alternately, take the 2-20 L micropipettor, put on appropriate tip and

1. Take a piece of parafilm* and place 5 individual 20 l drops of water on it.




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5. Transfer 8 L of colored fluid from the third drop into the fourth drop. Mix.

6. Transfer 4 L of colored fluid from the fourth drop into the fifth drop. Mix.

Alternately, take the 20-100 ml micropipettor, put on appropriate tip and

1. Take a piece of parafilm* and place 5 individual 100 ml drops of water on it.2. Transfer 50 L colored fluid from stock into the first drop. Mix.




Alternately, take the 200-1000 L micropipettor, put on appropriate tip and

1. Take a piece of parafilm* and place 5 individual 1000 l drops of water on it.





1000 l is equal to _______________ mL.

Measure and weigh 1000 L, 100 L and 20 L of distilled water transferred on parafilm using variousmicropipettors on a balance. One milliliter of water should weigh 1 g. Repeat 5 times, recording weight 5 times

on the table below.

Weight of drop in g

Trial 1 1000 L drop 100 L 20 L

2

3

4

5

Accuracy is an expression of how close a measurement instrument comes to the true or accepted value

1.0 mL of water should weigh 1.0 g. Look at your values. Comment on the accuracy of your micropipettor.

References:

Oklahoma City Community College Biotechnology Program (2009) Learning to Use a Micropipettor. Retrieved

15 June 2009 from http://www.occc.edu/ BBDiscovery/documents/Modules/ learn_micropip.htm

Oklahoma City Community College Biotechnology Program (2009) Additional Exercises in Using Micropipettors.

Retrieved 15 June 2009 from http://www.occc.edu/BBDiscovery/documents/

Modules/Micropipetting_Exercise.htm

SF Base (1994) Skill building Activity 1: Small volumes. Retrieved 15 June 2009 from

http://www.usc.edu/org/cosee-west/Jun07Resources/PipetteUsetraining.pdf


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Preparation of Chemical Reagents

Experiments in molecular biology (or any other field of science for that matter) require chemical reagents and

solutions. These must be correctly and accurately prepared for the experiment to succeed. Knowledge and

care in preparing solutions need to be mastered.

Calculating requirements for solution preparation

Solutions are mixtures of two or more substances with the relative amounts of each component being

specified. The amount of solute in a solvent is most often expressed in percentage, molarity or X.

1. Percentage, %

a. Percent weight by weight (%w/v): the number of grams of solute contained 100 mL of solution

Example: Prepare 10 mL of 1.0% solution of ammonium persulfate.

10 mL 0.01 = 0.1 g

Dissolve 0.1 g ammonium persulfate in 10 mL water.

b. Percent volume by volume (%v/v): the volume of solute contained in 100 mL of solution

Example: Prepare 250 mL of 70% isopropanol solution.

250 mL 0.7 = 175 mL

Mix 175 mL isopropanol with 75 mL water.

2. Molarity, M

Molarity is the number of moles of solute per liter of solution

M = mole/L, where M = molarity

L = volume of the solution in liters

mole = grams of solute (g) divided by molecular weight (g/mol)

Example: Prepare 500 ml of 0.5 M NaCl (NaCl is 58.55 g/mol). How many grams of NaCl do you need?

0.5 mol/L = X g

1

58.55 g/mol 0.5 L

X = 14. 6 g NaCl

Dissolve 14. 6 g NaCl powder in a final volume of 500 mL water.

3. Preparation of working solutions from concentrated stock solutions

Many buffers in molecular biology require the same components (Tris, EDTA, SDS, etc.) but often in varying

concentrations. To avoid having to make every buffer from scratch, it is useful to prepare several

concentrated stock solutions and dilute as needed.


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5. If the solution needs to be at a specific pH, check the pH meter with fresh buffer solutions and follow the

instructions for using a pH meter.

6. Make sure you label all prepared reagents properly, example:

7. Autoclave, if possible, at 121C, 15 psi for 20 minutes. Some solutions cannot be autoclaved; forexample, SDS. These should be filter-sterilized through a 0.22-m filter. When autoclaving liquids, do NOTtighten the caps to avoid pressure build up. Cover caps completely with aluminium foil. Media fo

bacterial cultures must be autoclaved the same day it is prepared, preferably within an hour or 2. Store

at room temperature and check for contamination prior to use by holding the bottle at eye level and

gently swirling it.

8. Solid media for bacterial plates can be prepared in advance, autoclaved, and stored in a bottle. When

needed, the agar can be melted in a microwave, any additional components, e.g., antibiotics, can beadded, and the plates can then be poured.

9. Concentrated solutions, e.g., 1M Tris-HCl pH = 8.0, 5M NaCl, can be used to make working stocks by

adding autoclaved double-distilled water in a sterile vessel to the appropriate amount of the

concentrated solution.

10.Glass and plasticware used for molecular biology must be scrupulously clean. Glassware should be

rinsed with distilled water and, if needed, autoclaved or baked at 150C for 1 hour. Plasticware, such as

pipettes and culture tubes, is often supplied sterile. Tubes made of polypropylene are turbid and

resistant to many chemicals, like phenol and chloroform; polycarbonate or polystyrene tubes are clear

and not resistant to many chemicals. Micropipette tips and microcentrifuge tubes should be

autoclaved before use. Boxes and jars of tips and tubes should be dried overnight in a drying oven

after autoclaving prior to use.

Reference:

Harisha, S. 2007. Biotechnology Procedures and Experiments Handbook. USA/India: Infinity Science Press LLC.


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Commonly used media and stock solutions

MEDIA

Super Broth (per liter)

Bacto-tryptone 10 g

Bacto-yeast extract 5 g

Glycerol 6.3 g

K2HPO4 12.5 g

KH2PO4 3.8 g

Autoclave.

LB (Luria Bertani) Medium (per liter)

Bacto-tryptone 10 g

Bacto-yeast extract 5 g

NaCl 10 gAutoclave. Where indicated add ampicillin to 50g/mL after autoclaved solution has cooled to 50oC.

LB plates with ampicillin, (per liter)

Add 15 g agar to 1 liter of LB medium. Autoclave. Allow the medium to cool to 55oC before adding ampicillin

(50 g/mL final concentration). Pour 30 - 35 mL of medium into 100 mm petri dishes. Let the agar harden. Storeat room temperature (for 1 week) or at 4oC (for 1 month).

ANTIBIOTICS

Ampicillin

Prepare a 50 mg/mL solution of ampicillin in H2O. Stock solutions of antibiotics dissolved in H2O should be

sterilized by filtration through a 0.22-micron filter. Store solutions in light-tight containers at -20oC.

STOCK SOLUTIONS

10% Sodium dodecyl sulfate (SDS)

Dissolve 10 g of electrophoresis-grade SDS in 90 mL of H2

O. Heat to 68o

C to assist dissolution. . Adjust volume to100 mL. CAUTION: wear a mask when weighing SDS. There is no need to sterilize 10% SDS.

1 M Tris

Dissolve 12.1 g Tris base in 80 mL of H2O Adjust the pH to the desired value by adding concentrated HCl:

pH 7.4: ~ 7.0 mL

pH 8.0: ~ 4.2 mL

pH 9.0: ~ 0.8 mL

Allow the solution to cool to room temperature before making the final adjustments to the pH. Make up the

volume of the solution to 100 mL. Dispense into aliquots and sterilize by autoclaving.

*NOTE: if you are using powdered Tris-HCl, check the pH before adjusting with additional HCl.

0.5 M EDTA (pH 8.0)

Add 186.1 g of disodium ethylene diamine tetraacetate-2 H2O to 800 mL of H2O Stir vigorously on a magneticstirrer Adjust the pH to 8.0 with NaOH (~20 g of NaOH pellets). Dispense into aliquots and sterilize by

autoclaving.

2M Glucose

Dissolve 18 g L-glucose (Dextrose) in 45 mL of H2O. Adjust volume to 100 mL. Filter sterilize.

10N NaOH


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Dissolve 20 g NaOH pellets in 40 mL of H2O. Adjust volume to 50 mL. Sterilize by autoclaving.

1M KCl

Dissolve 3.73 g KCl in 40 mL of H2O. Adjust volume to 50 mL. Sterilize by autoclaving.

1M NaCl

Dissolve 2.92 g NaCl in 40 mL of H2O. Adjust volume to 50 mL. Sterilize by autoclaving.

5 M Potassium acetate

To 60 mL of 5 M potassium acetate add 11.5 mL of glacial acetic acid and 28.5 mL of H2O. The resulting

solution is 3 M with respect to potassium and 5 M with respect to acetate. Sterilize by autoclaving.

Ethidium bromide 10 mg/mL

Add 1 g of ethidium bromide to 100 mL of H2O. Stir on a magnetic stirrer for several hours to ensure that the dye

has dissolved. Wrap the container in aluminum foil or transfer to a dark bottle and store at 4oC. CAUTION: EtBr is

a powerful mutagen. Wear gloves and a mask when weighing it out.

50 X TAE (Tris-Acetate) buffer, (per liter)

Tris base 242 g

Glacial acetic acid 57.1 mL0.5 M EDTA (pH 8.0) 100 mL

TE (sterile)

pH 8.0: 10 mM Tris-Cl (pH 8.0)

1 mM EDTA (pH 8.0)

TE (sterile)

pH 8.0: 10 mM Tris-Cl (pH 8.0)

0.1 mM EDTA (pH 8.0)

Gel loading buffer, Type III

6X buffer: 0.25% bromophenol blue

0.25% xylene cyanol

30% glycerol in H2O.

Store at 4oC.

Reference:

Sambrook, J, EF Fritsch and T Maniatis. 1989. Molecular Cloning: A Laboratory Manual. 2nd Edition. Cold Spring

Harbor Laboratory Press NY.

00 Pre-Laboratory Notes

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