Antelope Valley College

Introduction to Biotechnology Lab

Fall 2019

2

THE METRIC SYSTEM 3

BASIC LABORATORY SKILLS: PIPETTING 5

BASIC LABORATORY SKILLS: MEDIA AND SOLUTION PREPARATION 7

RESTRICTION ENZYME DIGESTION 10

GEL ELECTROPHORESIS 12

DNA EXTRACTION FROM AGAROSE GEL 14

PLASMID PREPARATION BY ALKALINE LYSIS 16

DOUBLE RESTRICTION ENZYME DIGESTION AND LIGATION 18

TRANSFORMATION OF CHEMICALLY-COMPETENT E. COLI 21

TRANSFORMATION OF ELECTROCOMPETENT E. COLI 23

BLUE/WHITE SCREENING FOR TRANSFORMANTS 25

DROSOPHILA GENOTYPE ANALYSIS USING PCR 27

MITOCHONDRIAL DNA ANALYSIS USING PCR 32

DETERMINATION OF PROTEIN MOLECULAR WEIGHT USING SDS-PAGE 36

BRADFORD PROTEIN ASSAY 40

3

The Metric System

Introduction

The metric system is a system of measurement that is based on multiples of 10. It is a

system that has been internationally agreed upon. It is synonymous with the International

System of Units (SI) and is used exclusively in almost every country in the world. The

simplicity of the metric system lies in the fact that it is based on powers of ten. This makes

converting between units of measure much simpler since there are no conversion values to

memorize (it’s just a matter of moving decimal places). In order to do metric system

conversions, two things do have to be known, though. First, one must know the prefixes that are

associated with the values at each level (for example, deci- is the prefix for the value 10-1 or

0.1). Second, one must also know in which direction the decimal will move in making

conversions (which is simple if you understand that moving the decimal to the right makes a

large number).

In the sciences, understanding math is crucial since it is what leads to the quantitative nature of

science. Understanding the metric system is crucial because it is the system used in the sciences,

worldwide. This gives everyone a chance to collaborate with people from all parts of the globe

using a “universal language”.

Materials

• Centimeter ruler

• Meter stick

• Electronic scale

• 200g calibration standards

• Graduated cylinders (10 & 100 mL)

• Beaker

Methods

A. Measuring length

1. Using the appropriate measuring tool, measure the height of your partner(s) in

centimeters. Convert all measurements to meters, millimeters, micrometers, and

nanometers. Express all measurements in scientific notation.

2. Using the appropriate measuring tool, measure the length of the right arm of your

partner(s) in centimeters. You will measure the arm from under the arm pit to the

longest fingertip. Convert all measurements to meters, millimeters, micrometers,

and nanometers. Express all measurements in scientific notation.

3. Using the appropriate measuring tool, measure the length of the right index finger

of your partner(s) in centimeters. You will measure the finger from the middle of

the largest knuckle (metacarpophalangeal joint) to the fleshy tip of the finger.

Convert all measurements to meters, millimeters, micrometers, and nanometers.

Express all measurements in scientific notation.

B. Measuring mass

1. Record the mass of the calibration weight standards given to ensure proper scale

calibration.

4

2. Using the electronic scale, measure each one of the various materials provided.

C. Measuring volume

1. Weigh the beaker that has been provided to you.

2. Using the graduated cylinders provided, measure out 5 mL of water into the

beaker.

3. Weigh the (filled) beaker and determine the weight of the water.

4. Record all data collected.

5

Basic Laboratory Skills: Pipetting

Introduction

In the biological sciences, it is crucial to be able to accurately and reproducibly measure

and transfer small volumes of liquids in order to obtain useful results. For volumes less than 1

ml, the most common (and preferred) method for measuring volumes involves the use of a

device known as a micropipette.

Diagram 2a shows an example of a micropipette that will be used in this class. [The devices you

use may not look exactly like the one shown. The micropipette used in this course come in three

different types: P1000, P200, and P20.]

Note that the minimum and maximum volume in microliters is marked on each micropipette.

Because of this, make sure that you are using the correct micropipette for the volume you need.

The micropipette needs to be set for the volume you need by looking in the “volume window”,

and, if necessary, turning the “volume control knob” until the micropipette displays the correct

volume. Before using the micropipette, always check that the desired volume is set. Never

assume that the volume will always be set for you. Do not attempt to set micropipette for

volumes larger than their maximum, or for volumes less than zero; doing so will damage

the micropipette.

Despite the obvious usefulness of micropipettes, they do carry with them a disadvantage.

If used improperly or even as a result of overuse, micropipettes may lose their calibration. This

will lead to improper volumes being pipetted. Because of this, there is a need to ensure that

calibration of the micropipettes is checked from time to time. Verifying the calibration of the

micropipettes will save the user time, material, and mistake-related frustration!

How to use the micropipette

All micropipettes use disposable tips. Do not pipet liquids without using the appropriate

tip, because this will contaminate the micropipette and will damage it. When attaching the tip,

make sure that the tip is properly seated on the end of the micropipette. Try depressing the

plunger. As the plunger depresses, you will feel first “stop”. If you continue pushing, you will

find a point where the plunger no longer moves downward (the second “stop”).

In order to use the micropipette, depress the plunger to the first “stop”, place the micropipette tip

into the liquid, and in a slow, controlled manner, allow the plunger to move upwards. Always

keep your thumb on the plunger; never just let the plunger go. Letting the plunger go will cause

the liquid to either splatter within the tip (leading to inaccurate volumes) or causing some of the

liquid enter the micropipette (contaminating it).

Take the micropipette (carrying the liquid in the tip) to the container to which you wish to

add liquid. In a continuous and steady manner, depress the plunger to the first, and then to the

second stop. If you watch carefully, you will note that depressing to the second stop expels all of

the liquid from the tip. In most cases, this will be true for most aqueous solutions. Sometimes,

however, it may be difficult to get all of the liquid out of the tip. In order to remedy this, it is

6

best to “wet” the tip, by pipetting the solution once, expelling it, and then taking up the liquid a

second time.

Materials

• Micropipette

• Micropipette tips

• Beakers

• Water

• Microcentrifuge tubes

• Erlenmeyer flask

Methods

A. Ensuring proper calibration

a. Obtain and record the weight of a beaker.

b. Using the proper micropipette, deliver 250 µL of water to the beaker.

c. Weigh the beaker and determine the weight of the water.

d. Verify that the micropipette is delivering the proper amount.

e. Repeat this process twice more for the same micropipette.

f. Repeat all these steps for

B. Practicing proper technique

a. Using proper technique (as demonstrated by the instructor), perform the

following procedure:

i. Obtain and record the weight of a microcentrifuge tube.

ii. Take 10 µL of water from a beaker to a microcentrifuge tube.

iii. Add 25 µL of water to the same microcentrifuge tube.

iv. Using a new tip, add 15 µL of dye to the same microcentrifuge tube.

v. Add 200 µL of water to the same microcentrifuge tube and mix all the

contents of the tube well.

vi. Now, take 250 µL out of the tube and discard.

vii. Weigh the tube and compare it to the original weight.

7

Basic Laboratory Skills: Media and Solution Preparation

Introduction

One of the most important skills in a biotechnology laboratory is media and solution

preparation. Arguably, there is nothing more reassuring to a researcher than knowing exactly

how the media and stock solutions were made. In molecular cloning experiments, there are two

media types that are usually made: selective media and enrichment media.

Selective media can be defined as one that either promotes the growth of an organism or

prevents the growth of others. Most of the selective media that is used in a biotechnology lab

will employ the use of antibiotics. In many cases, the media can also have a differential

component added. Differential media may reveal biochemical or physiological traits that the

organisms have. Say, for instance, that you introduced a gene that gives a bacterium that ability

to break down a substance that turns yellow once it is cleaved. If that material was introduced

into the medium and there was a color change, you would verify that you have the right

organism. What if there was no color change, however? That could tell you that you may have

the wrong organism or that you were unsuccessful in introducing the gene you wanted.

Enrichment media is designed to grow organisms. In many cases, you may see the term

“recovery medium”. These types of media contain all the requirements necessary for growth and

is used to ensure that the organisms that you are working with are thriving. Typically, Luria-

Bertani or Lysogeny Broth (LB) medium is used or, in some cases, Super Optimal Catabolite

(S.O.C.) medium will be used to grow (or promote recovery of) the cells used in these

experiments.

It is also crucial to understand how to make the different solutions that are found in a

biotechnology lab. Many of these solutions will be used to aid in the isolation of DNA or the

purification of proteins. Ensuring that the concentration of these solutions is as exact as possible

is also crucial. Using dilution formulas like C1V1=C2V2 and/or dimensional analysis are tools

that can be employed to determine the proper quantities

Materials

• DI water

• Screw-capped bottles

• Beakers

• Magnetic stir bars

• Sodium dodecyl sulfate (SDS) a.k.a. sodium lauryl sulfate

• Glucose

• Potassium acetate

• Tris base

• Concentrated HCl

8

• NaOH

• Plastic bottle/container

A. Making stock solutions

a. The following stock solutions need to be made:

i. 100 ml of 1 M glucose stock (filter-sterilized)

ii. 100 ml of 0.5 M Ethylenediaminetetraacetic Acid (EDTA), pH 8.0

iii. 100 ml of 10% SDS (autoclave)

iv. 100 ml of 10 N NaOH

v. 100 ml of 5 M potassium acetate (M.W.= 98.14) (autoclave)

vi. 100 ml of 1 M Tris-Cl

B. Making growth media

a. The following growth media need to be made:

i. 500 ml LB agar

ii. 250 ml LB broth

iii. Both media need to be autoclaved.

C. Recipes

a. To prepare EDTA at 0.5 M (pH 8.0): Add 186.1 g of disodium EDTA•2H2O to

800 mL of H2O. Stir vigorously on a magnetic stirrer. Adjust the pH to 8.0 with

NaOH (~20 g of NaOH pellets). Dispense into aliquots and sterilize by

autoclaving. The disodium salt of EDTA will not go into solution until the pH of

the solution is adjusted to ~8.0 by the addition of NaOH. Adjust the volume to 1

L using H2O if necessary.

b. To prepare a 1 M solution, dissolve 121.1 g of Tris base in 800 mL of H2O.

Adjust the pH to the desired value by adding concentrated HCl.

pH HCl

7.4 70 mL

7.6 60 mL

8.0 42 mL

Allow the solution to cool to room temperature before making final adjustments

to the pH. Adjust the volume of the solution to 1 L with H2O. Dispense into

aliquots and sterilize by autoclaving.

c. The preparation of 10 N NaOH involves a highly exothermic reaction, which can

cause breakage of glass containers. Prepare this solution with extreme care in

plastic beakers. As an added precaution, place the beaker on ice. When the pellets

9

have dissolved completely, adjust the volume to 1 L with H2O. Store the solution

in a plastic container at room temperature. Sterilization is not necessary

d. The following is the recipe for LB broth (per liter):

e. In order to make LB agar, the final concentration of agar needs to be 1.5%.

Tryptone 10 g

NaCl 5 g

Yeast extract 5 g

10

Restriction Enzyme Digestion

Introduction

Restriction enzymes are extremely important tools in biotechnology. Restriction

enzymes cleave DNA only at specific sequences. Different bacteria contain a large variety of

restriction enzymes, and enzymes with specificity for large numbers of sequences are

commercially available. In order to decide which restriction enzymes to use for a particular

experiment, you will need to know the DNA sequence of the gene of interest as well as the

plasmid map of the vector you are using. There are a number of resources to aid in the process

of obtaining sequence information, including the NCBI website.

In order to prepare for construction of a new plasmid, we will first verify the identity of

the plasmid we are using. We will do this using a technique called restriction mapping. Here we

will employ the use of a plasmid map or other resources to determine what restriction enzymes to

use and predict the size of the resulting fragment(s).

Restriction enzymes must cleave both strands of the double stranded DNA. They can do

this in a number of ways. Some restriction enzymes cleave both strands at the same location,

resulting in a “blunt” end. Other restriction enzymes cleave at different locations on the different

strands, leaving short “overhangs” of single stranded DNA also known as “sticky ends”. These

staggered end cuts are desired over their blunt end counterparts because it may facilitate ligation

of foreign segments of DNA into the plasmid vector. In order to prepare for ligation, then, the

plasmid and the gene of interest (foreign DNA) must be digested with two different restriction

enzymes.

A potential problem with using multiple restriction enzymes is that each one is active in

specific buffers; the best buffer for one enzyme may not be the best one for another enzyme.

However, recently, there have been companies that have engineered buffers that can host a wider

range of restriction enzymes. Traditionally, however, it is necessary to run one reaction,

deactivate the enzyme or perform a DNA purification, and then run the other reaction. In other

cases, there may be a complementary pair of enzymes that share a buffer and a double digest

may be performed.

Restriction enzymes should be kept on ice at all times, and solutions containing them

should be buffered at the proper pH. The concentration of a restriction enzyme is usually

expressed in enzyme units per volume. One unit of restriction enzyme is defined as the amount

of enzyme needed to digest 1 μg of DNA in 1 hour.

Materials

• Megapure water

• Lambda DNA/EcoRI + HindIII Marker

• Lambda DNA

• Plasmid DNA

11

• FastDigest Restriction Enzymes

o We may use:

▪ EcoRI

▪ HindIII

▪ XbaI

▪ XhoI

• FastDigest Green Buffer

o Proprietary buffer from Thermo Scientfic

▪ Universal buffer with optimal pH, glycerol concentration, and salt

concentration

o Supports 100% activity of all FastDigest restriction enzymes

o No star activity

Methods

A. Restriction enzyme digest

a. Set up each digestion as follows:

Water 15 µL

10X Fast Digest Buffer or 10X

Fast Digest Green Buffer 2 µL

DNA 2 µL

FastDigest enzyme 1 µL

b. Mix all components well and spin down.

c. Incubate the reaction for 5 minutes at 37C.

d. View results of restriction enzyme digest using gel electrophoresis or place tubes

in -20C for future analysis

12

Gel Electrophoresis

Introduction

The purpose of agarose gels is to allow for the estimation of DNA fragment size. This

can be useful for verifying that a fragment of the correct size was produced in a polymerase

chain reaction or to assess the size of restriction enzyme digestion fragments. It is also useful to

run plasmid DNA on a gel in order to estimate both the concentration of the DNA and the quality

of the preparation (i.e. to look for possible contaminants or excessive fragmentation of the

plasmid DNA).

Remember that DNA is a negatively charged molecule due to their phosphate backbone.

Agarose forms a low-density matrix and is therefore suited to running the large DNA molecules.

Various concentrations of agarose can be used to separate different sized DNA: 0.75% for DNA

> 3 kb, 2% or 3% for 50 – 400 base pairs (bp), and 1% for DNA between these ranges. In our

lab, however, we will run 0.7% agarose gels. Agarose gels are usually run in either TBE (Tris-

borate-EDTA) or TAE (Tris-acetate- EDTA) buffer (as it is in our lab). The agarose comprises a

very small percentage of the gel while the remainder is buffer and because of this, the gel is

merely submerged in the buffer, with the electrical current running through both the buffer and

the gel. This is why agarose gel electrophoresis is also known as “submarine electrophoresis”.

In order to correctly determine the size of the resulting DNA/fragments, it is necessary to

also load and run molecular weight size marker/ladder. These “ladders” can be DNA, RNA, or

protein (depending on the electrophoresis application used) and, in the case of DNA at least,

some type of DNA that is digested with a restriction enzyme or combination of enzymes. The

molecular weight of the nucleic acid/protein is inversely proportional to the migration rate

through some type of gel matrix (again, depending on the application). Since the ladder provides

fragments of different sizes, they can be used to estimate the size of the results of gel

electrophoresis.

Materials

• Agarose

• 50X TAE buffer

• Gel electrophoresis chamber

• Power supply

• Gel casting material

o Gel tray

o Rubber dams

o Plastic comb

Methods

A. Agarose gel preparation

a. Make 60 ml of 0.8% agarose

13

i. Make 1L of 1X TAE buffer using 50X stock

ii. Dissolve agarose in 1X TAE buffer

iii. Heat agarose and buffer in microwave for 2 minutes

iv. Use hot gloves to remove flask from microwave and begin to cool in a

50C water bath

b. Add 6 µL of SYBR Safe stain to the warm agarose and mix well.

c. Pour warm agarose into gel mold (with rubber blocks and comb installed)

d. When the gel is solidified, remove the rubber blocks. The gel is now ready to be

placed in the gel electrophoresis chamber.

B. Ladder/Marker Preparation

a. Heat the ladder for 5 minutes at 65C and cool on ice for 3 minutes. Prepare the

DNA ladder as follows:

Water 16 µL

6X DNA Loading Dye 3 µL

DNA ladder 1 µL

C. Sample Preparation

a. You will run the samples you prepared last week or you may prepare uncut

plasmid or DNA samples using the following recipe:

Water 15 µL

6X DNA Loading Dye 3 µL

DNA sample 2 µL

D. Gel electrophoresis

a. Here is an example of lane assignments you can use:

i. Empty

ii. Plasmid DNA (undigested)

iii. Digested plasmid DNA

iv. Lambda DNA EcoRI/HindIII ladder

v. Lambda DNA (undigested)

vi. Lambda DNA Digest I

vii. Lambda DNA Digest II

viii. Lambda DNA EcoRI/HindIII ladder

b. Run the gel at ~60 V for about 90 minutes.

c. View the gel using a transilluminator or gel visualization device.

14

DNA Extraction from Agarose Gel

Introduction

In many cases, it is desirable to extract DNA from an agarose gel. This allows you to

isolate and then purify DNA fragment based on their size (length in base pairs). While this can

be done using methods presented by researchers (including Sun et. al.), this can be accomplished

using commercially available kits. Note: The following protocol is an excerpt from the

ThermoScientific GeneJet Gel Extraction and DNA Cleanup Kit #K0832

Methods

1. Excise up to 200 mg gel slice containing the DNA fragment using a clean scalpel or razor

blade. Cut as close to the DNA as possible to minimize gel volume. Place the gel slice

into a 1.5 ml tube. Note: If the purified fragment will be used for cloning reactions,

avoid damaging the DNA through UV light exposure. Minimize UV exposure to a few

seconds or keep the gel slice on a glass or plastic plate during UV illumination.

2. Add 200 µL of Extraction Buffer. Mix thoroughly by pipetting.

3. Incubate the gel mixture at 50-58C for 10 minutes or until the gel slice is completely

dissolved. Mix the tube by inversion every few minutes to facilitate the melting process.

Ensure that the gel is completely dissolved. Note: For >1% agarose gels, prolong the

incubation time to 15 minutes.

4. Add 200 µL of ethanol (96-100%). Mix thoroughly by pipetting.

5. Transfer the mixture to the DNA Purification Micro Column preassembled with a

collection tube. Centrifuge the column for 30-60 seconds at 14,000 x g. Discard the

flow through. Place the DNA Purification Micro Column back into the collection tube.

Note:

a. If DNA fragment is > 10 kb centrifuge the column for 2 minutes at 14,000 x g.

b. Close the bag with DNA Purification Micro Columns tightly after each use.

6. Add 200 µL of Prewash Buffer (supplemented with ethanol) to the DNA Purification

Micro Column and centrifuge for 30-60 seconds at 14,000 x g. Discard the flow-through

and place the purification column back into the collection tube.

Note: If DNA fragment is > 10 kb centrifuge the column for 1-2 minutes at 14,000 x g.

7. Add 700 µL of Wash Buffer (supplemented with ethanol) to the DNA Purification

Micro Column and centrifuge for 30-60 seconds at 14,000 x g. Discard the flow-through

and place the purification column back into the collection tube.

Note: If DNA fragment is > 10 kb centrifuge the column for 1-2 minutes at 14,000 x g.

8. Repeat step 7.

9. Centrifuge the empty DNA Purification Micro Column for an additional 1 minute at

14,000 x g to completely remove residual Wash Buffer. Note: This step is essential to

avoid residual ethanol in the purified DNA solution. The presence of ethanol in the DNA

sample may inhibit downstream enzymatic reactions.

15

10. Transfer the DNA Purification Micro Column into a clean 1.5 ml microcentrifuge tube.

11. Add 10 µL of Elution Buffer to the DNA Purification Micro Column. Centrifuge for 1

minute at 14,000 x g to elute DNA.

Note:

a. If DNA fragment is > 10 kb the elution volume should be increased between 15-

20 µL to get optimal DNA yield. Elution volume less than 10 µL is not

recommended.

b. Lower volume of Elution buffer for DNA Micro Kit can be used (6-10 µL) in

order to concentrate eluted DNA. Please notice that < 10 µL elution volume

decreases DNA yield.

12. Discard the purification column and store the purified DNA at -20C.

16

Plasmid Preparation by Alkaline Lysis

Introduction

Recall that plasmids are extrachromosomal DNA and may contain genes that will give

bacteria the ability to have antibiotic resistance, the ability to produce pili, and the ability to

metabolize sugars or material that they normally could not. Researchers have engineered

plasmids to contain a polylinker or multiple cloning site (MCS) which is a segment of the

plasmid that contains several restriction enzyme (RE) sites. The RE sites within the MCS occur

usually are unique sites, occurring only once in the plasmid to ensure that researchers can insert

foreign DNA within the site.

Materials

• Alkaline lysis solution I

• Alkaline lysis solution II (prepared fresh and used at room temperature)

• Alkaline lysis solution III

• Microcentrifuge tube

• Bacterial culture

Methods

A. Preparation of cells

a. Inoculate 2 ml of LB (with appropriate antibiotic) with a single (plasmid-

containing) bacterial colony.

b. Incubate the culture overnight at 37C in an incubator shaker.

c. After the culture has grown, take 1.5 ml and centrifuge at maximum speed for 30

seconds at 4C.

d. Remove the supernatant (the culture medium), leaving the bacterial pellet as dry

as possible. It is very important to ensure that the pellet is as dry as possible.

B. Lysis of cells

a. Resuspend the bacterial pellet in 100 µL of ice-cold alkaline lysis solution I by

vortexing vigorously.

b. Add 200 µL of freshly prepared alkaline lysis solution II. Close the tube tightly

and mix the tube by inverting it rapidly 5 times. Do not vortex! Store the tube

on ice.

c. Add 150 µL of ice-cold alkaline lysis solution III. Close the tube tightly and mix

the tube by inverting it several times. Do not vortex! Store the tube on ice for 3-

5 minutes.

d. Centrifuge the bacterial lysate at maximum speed for 5 minutes at 4C. Transfer

the resulting supernatant to a fresh tube.

17

C. Recovery of plasmid DNA

a. In order to precipitate the nucleic acid, add two volumes of room temperature

ethanol to the supernatant. Mix the solution by vortexing and let it stand for 2

minutes at room temperature.

b. Collect the nucleic acids by centrifugation at maximum speed for 5 minutes at

4C.

c. Remove the supernatant by aspirating gently with a pipette. Remove as much of

the supernatant as possible.

d. Once dry, add 1 ml of ice-cold 70% ethanol to the pellet and invert the closed

tube several times. Centrifuge the tube at maximum speed for 2 minutes.

e. Remove the supernatant by aspirating gently with a pipette. Remove as much of

the supernatant as possible.

f. Remove any beads of ethanol that are on the sides of the tube. Store the open

tube at room temperature until all the ethanol has evaporated (about 10-15

minutes).

g. Dissolve the DNA in 50 µL of TE (pH=8.0). Vortex the solution gently for a few

seconds. Store the DNA at -20C.

18

Double Restriction Enzyme Digestion and Ligation

Introduction

The process of connecting two pieces of DNA together is called ligation. Ligase is the

enzyme used to catalyze the reaction of binding two pieces of DNA together. The ligase used

most often in recombinant DNA technology is derived from the T4 bacteriophage. Much like

ligase found in cells, T4 DNA ligase uses ATP to supply the energy necessary for the reaction.

In order to be able to ligate a DNA molecule, there needs to be DNA with a 5 ́-phosphate

group and free 3 ́-hydroxyl groups. In other words, we must use DNA that has been digested

with restriction endonucleases. As you may recall, products from restriction enzyme digests can

be blunt ended (i.e. all of the bases are paired with bases from the opposite strand) while others

leave “overhangs” or “sticky ends”, which are short stretches of single stranded DNA at either

the 5 ́ or 3 ́ end.

Materials

• Plasmid DNA

• Gene of interest (foreign DNA)

• Water

• FastDigest Restriction Enzymes

• Water bath

• GeneJet Gel Extraction and DNA Cleanup Micro Kit

• ThermoFisher Rapid DNA Ligation Kit

o T4 DNA Ligase, 5u/µL

o 5X Rapid Ligation Buffer

o Nuclease free water

• Microcentrifuge tubes

• Double-digested plasmid vector

• Double-digested insert DNA (or double-digested PCR product)

• Room temperature (22C) condition

Methods

A. Restriction enzyme digest

a. Set up each double digestion as follows:

Water 14 µL

10X Fast Digest Green Buffer 2 µL

DNA 2 µL

FastDigest enzyme (each) 1 µL

b. Mix all components well and spin down.

19

c. Incubate the reaction for 20 minutes at 37C.

d. View results of restriction enzyme digest using gel electrophoresis.

B. DNA purification

a. Purify the digested plasmid and gene of interest using the following protocol from

the GeneJet Gel Extraction and DNA Cleanup Micro Kit:

i. Adjust the volume of the reaction mixture to 200 μL with nuclease-free

water or TE buffer.

ii. Add 100 μL of Binding Buffer. Mix by pipetting.

iii. Add 300 μL of ethanol (96-100%) and mix by pipetting.

iv. Transfer the mixture to the DNA Purification Micro Column preassembled

with a collection tube. Centrifuge the column for 30-60 seconds at 14,000

× g. Discard the flow-through. Place the DNA Purification Micro Column

back into the collection tube.

1. If DNA fragment is ≥ 10 kb centrifuge the column for 2 minutes at

14,000 × g.

2. Close the bag with DNA Purification Micro Columns tightly after

each use!

v. Add 700 μL of Wash Buffer to the DNA Purification Micro Column and

centrifuge for 30-60 seconds at 14,000 × g. Discard the flow-through and

place the purification column back into the collection tube.

1. If DNA fragment is ≥ 10 kb centrifuge the column for 2 minutes at

14,000 × g.

vi. Repeat step 5.

vii. Centrifuge the empty DNA Purification Micro Column for an additional 1

minute at 14,000 × g to completely remove residual Wash Buffer.

1. This step is essential to avoid residual ethanol in the purified DNA

solution. The presence of ethanol in the DNA sample may inhibit

downstream enzymatic reactions.

viii. Transfer the DNA Purification Micro Column into a clean 1.5 mL

microcentrifuge tube.

ix. Add 10 μL of Elution Buffer to the center of the DNA Purification Micro

Column membrane. Centrifuge for 1 minute at 14,000 × g to elute DNA.

1. If DNA fragment is ≥ 10 kb the elution volume should be

increased to 15-20 μL to get optimal DNA yield.

2. Lower volume of Elution Buffer for DNA Micro Kit can be used

(6-10 μL) in order to concentrate eluted DNA. Please notice that <

10 μL elution volume slightly decreases DNA yield.

3. Double the elution volume or perform two elution cycles when

purifying larger amounts of DNA (for example > 5 μg).

20

x. Discard the purification column and store the purified DNA at -20°C.

C. Rapid ligation

a. Label a microcentrifuge tube appropriately.

b. In the tube, add:

Double-digested plasmid DNA 10-100ng

Double-digested insert DNA 3:1 insert to vector ratio

5X Rapid Ligation Buffer 4 µL

T4 DNA Ligase 1 µL

Nuclease-free water Bring to 20 µL volume

c. Vortex the tube and then briefly spin down the reaction mixture.

d. Incubate the mixture at 22C for 5 minutes.

e. Confirm successful ligation by performing gel electrophoresis

f. Proceed to transformation or store the reaction mixture at 0-4C.

21

Transformation of Chemically-Competent E. coli

Introduction

In order to produce more plasmids, the insertion of plasmid DNA into bacterial cells is

necessary. In general, bacteria do not naturally take up DNA from their environment. In order

to improve the probability that the bacteria will internalize the plasmid DNA, it is necessary to

make the cells “competent” to take in the plasmid DNA. Competent cells are significantly more

fragile than normal bacteria. Vortexing the cells, heating the cells above 42°C or to 42°C for

prolonged periods, or exposure of the cells to any of a number of other even mildly abusive

treatments may kill them. If done correctly, the cells will break open long enough to take in any

plasmid DNA that is in the tube. The cells are then allowed to “heal”. This process is called

transformation.

Materials

• 37°C shaking and non-shaking incubator

• LB agar plates with 100 µg/ml of ampicillin

• 42°C water bath

• Ice Bucket

• Foam test tube rack

• One Shot Top 10 Competent Cells (CaCl2-competent E. coli)

• Plasmid DNA

Methods

A. Preparation

a. Ensure water bath is at 42°C.

b. Make sure S.O.C. medium is at room temperature.

c. Spread X-gal onto LB Amp 100 plates.

B. Transformation

a. Centrifuge tube(s) containing the ligation reaction and place the tube on ice.

b. Obtain a vial of competent cells and place it on ice to thaw (this may have already

been done for you).

c. Aseptically transfer 1-5 µL of your ligation reaction into the competent cells.

Mix by tapping. DO NOT MIX BY PIPETTING.

d. Incubate the tubes on ice for 30 minutes.

e. Transfer the test tube into the 42°C water bath and incubate for EXACTLY 30

seconds. This heat shock step is extremely time-sensitive and must be performed

very accurately. DO NOT MIX OR SHAKE.

f. Immediately put the test tube back on ice.

g. Aseptically add 250 µL of S.O.C. to the tube.

22

h. Tape the tube on its side to the inside of a shaking incubator. Shake the vials at

37°C for 1 hour at 225 rpm.

i. Spread 20-200 µL of the transformation vial onto the appropriately labeled agar

plate.

j. Using a sterile loop, perform a “lawn” streak of the bacterial culture.

k. Allow the plates to sit, lid side up, for a few minutes before inverting the plates.

l. Tape together the plates and incubate them, lid side down, at 37°C. These plates

will be incubated for 24 hours.

m. Colonies will be selected and analyzed in the next class period.

23

Transformation of Electrocompetent E. coli

Introduction

In order to produce more plasmids, the insertion of plasmid DNA into bacterial cells is

necessary. In general, bacteria do not naturally take up DNA from their environment. In order

to improve the probability that the bacteria will internalize the plasmid DNA, it is necessary to

make the cells “competent” to take in the plasmid DNA. Competent cells are significantly more

fragile than normal bacteria. Vortexing the cells, heating the cells above 42°C or to 42°C for

prolonged periods, or exposure of the cells to any of a number of other even mildly abusive

treatments may kill them. If done correctly, the cells will break open long enough to take in any

plasmid DNA that is in the tube. The cells are then allowed to “heal”. This process is called

transformation.

Materials

• 37°C shaking and non-shaking incubator

• LB agar plates with 100 µg/ml of ampicillin

• Ice Bucket

• One Shot Top 10 Electrocompetent Cells (electrocompetent E. coli)

• Electroporation cuvettes

• Electroporator

• Plasmid DNA

Methods

A. Preparation

a. Ensure water bath is at 42°C.

b. Make sure S.O.C. medium is at room temperature.

c. Spread X-gal onto LB Amp 100 plates.

B. Transformation

a. Thaw, on ice, one vial of electrocompetent cells for each transformation.

b. Add 1-2 μl of the DNA (10 pg to 100 ng) into a vial of electrocompetent cells and

mix gently.

c. Transfer the cells to the chilled electroporation cuvette on ice.

d. Electroporate the cells as per the manufacturer’s recommended protocol.

e. Aseptically add 250 μl of pre-warmed S.O.C. Medium to each vial.

f. Transfer the solution to a 15 ml snap-cap tube and shake for at least 1 hour at

37°C to allow expression of the antibiotic resistance gene.

g. Spread 10 to 150 μl from each transformation on a pre-warmed selective plate and

incubate overnight at 37°C. We recommend that you plate two different volumes

to ensure that at least one plate will have well-spaced colonies. For pUC19, dilute

24

the transformation mix 1:50 into LB Medium (e.g. remove 20 μl of the

transformation mix and add to 980 μl of LB Medium) and plate 20-100 μl.

h. Store the remaining transformation mix at +4°C. Additional cells may be plated

out the next day, if desired.

i. Invert the selective plate(s) and incubate at 37°C overnight.

j. Select colonies and analyze by plasmid isolation, PCR, or sequencing.

C. Electroporation Settings

Choose

Mode:

T HV

Set

Capacitance:

C Default setting is 50 μF in HV

Set

Resistance:

R R5 (129 ohms)

Chamber

Gap:

1 mm gap

Set

Charging

Voltage:

S 1.3 – 1.5KV

Desired

Field

Strength:

E 13.0 – 15.0 KV/cm

Desired

Pulse

Length:

t 5 – 6 msec

25

Blue/White Screening for Transformants

Introduction

It is important to be able to test whether or not transformation was successful. As it is,

transformation is usually an extremely inefficient process; in most cases, only a very small

fraction of cells actually takes in the DNA. If it were necessary to sort out the few cells that took

up DNA from the vast majority that did not, cloning experiments would be very difficult.

However, selection mechanisms make this fairly straightforward.

The selection mechanism most commonly used is antibiotic resistance. If the E. coli

strain used is not resistant to the antibiotic, but the plasmid DNA contains a gene coding for

resistance, then only the cells that have taken up the plasmid will be able to grow. Therefore, if

organisms show growth on media that contains that antibiotic, it demonstrates that the plasmid

was successfully taken in by the competent cells. It does not, however, confirm if the gene of

interest was inserted correctly into the MCS of the plasmid.

Researchers discovered that deleting a section from the lacZ gene (a mutation called

lacZΔM15) creates a non-functional β-galactosidase enzyme (the enzyme responsible for lactose

metabolism). Providing DNA encoding this section of amino acids (called the α-peptide) to a

lacZΔM15-mutant bacterial cell in trans complements the mutation allowing for a functional

enzyme. This process is called α-complementation. Researchers engineered an MCS into the α-

peptide and inserted it into a plasmid (like pUC19), creating an α-complementation cloning

vector. When a cloning experiment is successful and the insert (foreign) DNA is cloned into the

MCS of these plasmids, the α-peptide gets interrupted and thus will not complement the cell's β-

galactosidase mutation. If the reaction is unsuccessful, it leaves the α-peptide intact, and the cell

will have a functional β-galactosidase enzyme.

The organism will then be able to cleave lactose. The cleavage of lactose does not really

help researchers determine whether or not the α-peptide got interrupted. Therefore, we employ

the use of X-gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside), a colorimetric substrate

that will normally turn blue if the bacterial cells have an intact α-peptide (and a functional β-

galactosidase). If the cells turn white, however, the cells will have an interrupted α-peptide,

meaning successful insertion of the gene of interest. Isopropyl β-D-1-thiogalactopyranoside

(IPTG) is needed for induction of the lac operon and therefore a necessary component for the

cleavage (or lack thereof) of the X-gal substrate.

Materials

• LB agar plates with 100 µg/ml of ampicillin

• X-gal

• IPTG

• Cells from transformation

Methods

26

A. Blue-white screening

a. Spread 40 µL or appropriate amount of stock solution of X-gal and 10 µL of

IPTG solution on LB ampicillin plates using a sterile spreader.

b. Preferably, leave the plates to dry in laminar flow chamber with lids slightly

open.

c. Spread 10-100 µL of transformed E. coli cells onto the LB agar plates using

sterile spreader.

d. Incubate the plates at 37°C for 24 hours.

e. Blue and white colonies appear on the agar surface. Select the recombinant

cells in the white colonies to culture.

27

Drosophila Genotype Analysis Using PCR

Introduction

A model organism is any plant, animal or microorganism that allows us to study

fundamental questions in biology that may be hard to study directly in complex organisms like

humans. For almost a century, the fruit fly Drosophila melanogaster has been a valuable model

organism for research in genetics, developmental biology and evolutionary biology. In the early

1900’s, Thomas Hunt Morgan used the fruit fly to illustrate one of the core principles of modern

genetics – the linkage of a gene with a particular chromosomal location. For this groundbreaking

work in genetics, Morgan was awarded the Nobel Prize in Physiology or Medicine. Fruit flies

were initially chosen as a model organism because they have a simple genome, a fast generation

time, and they breed in large numbers. A new generation of fruit flies can be produced every

two weeks. True-breeding fruit flies are put together in a culture vial and allowed to mate.

Drosophila larvae will hatch twenty-four hours after the female fly deposits fertilized eggs on

special growth media. The larvae will then molt one, two, and four days after hatching,

increasing in size each time. After the third larval molt, the fly pupates for four days. After

metamorphosis, Drosophila emerges from the pupa as a winged adult with brick red eyes and

yellow-brown bodies with black rings on the abdomen. Adults are fertile after two days, which

means that these flies can quickly be used for further genetic studies.

Today, the entire Drosophila genome has been completely sequenced and several thousand

genetic mutants are available for study. This allows scientists to correlate changes at the DNA

level with changes in phenotype. Many genes found to be crucial in human health and

development are conserved in Drosophila. In fact, about 75% of the genes that cause dis- ease in

humans have homologs in the fruit fly. This makes the humble fruit fly an important model

system in today’s research laboratory. The wide array of Drosophila mutants available to

geneticists has been generated through many different means, including X-rays, mutagenic

chemicals and mobile genetic elements known as transposons. There are many different types of

transposons found in the Drosophila genome, including P-elements, Doc, copia and gypsy.

When transposons “jump” into relatively random sites throughout the Drosophila genome, they

can disrupt genes -- in fact, transposons account for over 50% of spontaneous mutations

identified in Drosophila!

The Doc element belongs to a group of retrotransposons known as long interspersed elements, or

LINEs, which are found in abundance in eukaryotic genomes. Like other retrotransposons, the

Doc element is believed to “hop” through the Drosophila genome using an RNA intermediate.

In brief, the Doc element is transcribed into mRNA using the fly’s native DNA polymerase. The

Doc mRNA codes for an enzyme called Reverse Transcriptase (RT), which can synthesize a

strand of DNA using an RNA template. After the RT mRNA is translate in the cytoplasm, the

protein is transported back into the cell’s nucleus, where it converts the Doc mRNA into DNA.

The newly synthesized DNA integrates at a new location in the genome, while the original copy

of Doc transposon remains at the same genomic location. For this reason, eukaryotic genomes

often contain thousands of copies of LINE retrotransposons.

28

Thomas Hunt Morgan used eye color mutants to discover the relationship between genes and

chromosomes. While sorting through his flies, he identified a single white-eyed male among the

red-eyed flies. This mutation, called white-1, was the first genetic mutant identified in

Drosophila. Morgan crossed the white-eyed fly with true-breeding red-eyed female flies. All of

the flies in the first filial generation (or F1) had red eyes, suggesting that white allele was

recessive to the red allele. To confirm this, Morgan crossed male and female flies from the F1

generation. The second generation (second filial, or F2) observed the classical Mendelian ratio

of three red-eyed flies for every one white-eyed fly. However, all of the female flies in the

second generation were red-eyed. Half of the males were red-eyed, while the rest had white eyes

like the original mutant. From this data, Morgan reasoned that the gene responsible for the white

color of the eye was located on the X-chromosome, providing the first evidence of a gene being

linked to a specific chromosome. Since white is a sex-linked gene, males will exhibit the white-

eyed phenotype with one copy of the white-1 mutant, as they have one X chromosome and one Y

chromosome. Females must have two copies of the recessive allele to see the white-eyed

phenotype. As such, crosses between white-eyed females and males will only produce white

eyed offspring. In contrast, crossing white-eyed females with wild-type males will result in

female offspring with red eyes and males with white eyes

Today, scientists have shown that eye color in Drosophila results from a combination of

pigments known as the pteridines and the ommochromes. Two separate enzymatic pathways

work in concert to synthesize the pigments from amino acid precursors. The pteridine pathway

converts the tryptophan into a bright red pigment and the ommochrome pathway converts the

guanine into a brownish pigment. The white gene codes for a special transporter protein that

moves the precursor amino acids into the developing eyes, allowing the pigments to be

synthesized. The white-1 mutation initially described by Morgan was caused by the insertion of

the 4.4 kilobase pair (kb) Doc transposon into the promoter region of the white gene. The large

transposon blocks transcription of the messenger RNA, meaning that the transporter protein

cannot be produced. Without this protein, neither of the pigments can be synthesized, so white-1

flies have white colored eyes. Because of this, white is considered to be the master eye color

gene.

In 1984, Dr. Kary Mullis revolutionized the field of molecular biology when he devised a simple

and elegant method to copy specific pieces of DNA. Recognizing that an initial step in DNA

replication in a cell’s nucleus is the binding of RNA primers, Mullis discovered that he could

replicate DNA in vitro using short, synthetic DNA primers and DNA Polymerase I.

Furthermore, because researchers can specify a primer’s sequence to target a specific gene, this

method allowed for the rapid amplification of a selected DNA sequence in vitro. For the

development of this technique, known today as the Polymerase Chain Reaction (or PCR), Mullis

was awarded the Nobel Prize in Chemistry in 1993.

Because of its ease of use and its ability to rapidly amplify DNA, PCR has become indispensable

in life sciences labs, replacing the time-intensive Southern blot as the method of choice to

characterize genetic differences at the molecular level. To perform PCR, purified double-

stranded DNA is mixed with primers (short synthetic DNA molecules that target DNA for

amplification), a thermostable DNA polymerase (Taq) and nucleotides. The mixture is heated to

94°C to denature the DNA duplex (i.e., unzip it into single strands). Next, the sample is cooled

29

to 45°C-60°C, allowing the primers to base pair with the target DNA sequence (called

“annealing”). Lastly, the temperature is raised to 72°C, the optimal temperature at which Taq

polymerase will extend the primer to synthesize a new strand of DNA. Each “PCR cycle”

(denaturation, annealing, extension) doubles the amount of the target DNA in less than five

minutes. In order to produce enough DNA for analysis, twenty to forty cycles may be required.

To simplify this process, a specialized machine, called a “thermal cycler” or a “PCR machine”,

was created to rapidly heat and cool the samples to the designated temperature for each of the

three steps. PCR is a simple, fast and reliable method to identify specific genetic mutations. For

example, wild type and white-1 flies can be distinguished using primers that flank the Doc

element present in the white promoter. After PCR amplification, DNA extracted from flies

without the Doc transposon will produce a 220 base pair (bp) amplicon, whereas DNA extracted

from white-1 flies will produce a 4.6 kb amplicon. However, since Taq polymerase is not

efficient at amplifying DNA fragments greater than 2 kb, this large fragment is not usually

produced. An unlinked gene coding for a wing protein is simultaneously amplified as a positive

control; this PCR product is 1000 bp.

Materials

• Thermo cycler

• Screw-capped tubes

• Microcentrifuge tube

• Ice Bucket

• Test tube rack

• Water bath

• PCR primers

• PCR Edvobead

o DNA polymerase

o Polymerase buffer

o dNTP mix

• Saline solution

Methods

A. Isolation of DNA from Drosophila

WARNING!

Use only screw-cap tubes when incubating in the water bath for DNA isolation. Do

not use snap-top tubes.

a. Label one 0.5 ml screw-cap tube with “wild” and a second 0.5 ml screw-cap tube

with “white”.

b. Anesthetize the fruit flies by placing the vials in the freezer for one to two

minutes.

c. Transfer one wild type fly from the vial to the tube labeled “wild”. Using a clean

pipet tip or toothpick, mash the fly in the tube (about 10 seconds).

30

d. Using a fresh pipet tip, add 100 μl of the DNA extraction buffer to the fly. Mix

the sample by pipetting up and down.

e. Place the “wild” tube on ice. Repeat steps 3-4 with the white-eyed flies using

fresh tubes and tips.

f. Incubate the samples in a 70° C water bath for 15 minutes.

g. Add 14 μl of potassium acetate to each sample and mix for 5 seconds.

h. Incubate on ice for 5 minutes.

i. Centrifuge the samples at maximum speed for 5 minutes.

j. Carefully transfer the supernatant to clean, labeled 0.5 ml snap-top

microcentrifuge tubes. Avoid the pellet at the bottom of the tubes. After removing

the supernatant, discard the tubes with the pellet.

k. Add 45 μl of room temperature isopropanol to each sample to precipitate the

DNA.

l. Centrifuge the samples at maximum speed for 5 minutes.

m. After centrifugation, a very small DNA pellet should be visible at the bottom of

the tube. With a marker, circle the location of the DNA pellet.

n. Carefully remove and discard the supernatant. Take care to avoid the DNA pellet

while removing the supernatant. Carefully wash the pellet with 20 μl of 70%

EtOH.

o. Centrifuge the tubes at maximum speed for 2 minutes.

p. Carefully remove and discard the supernatant. Allow the pellet to completely dry

(5-10 minutes)

q. Resuspend the DNA pellet in 25 μl of TE buffer.

r. Place tubes in ice

B. Amplification of the DNA

a. Label two PCR tubes "wild" and "white". If assigned control DNA, label a third

PCR tube “control”.

b. To each tube, add one PCR edvobead, 20 μl primer mix, and 5 μl extracted DNA

(either wild, white, or control DNA).

c. Mix the PCR samples gently. Make sure the PCR edvobeads are completely

dissolved in each tube.

d. Centrifuge to collect the samples at the bottom of the tubes.

e. Alternatively (and usually), a PCR master mix comprising of all the components

that are common to the reaction can be made up. Master mixes can aid in less

contamination being introduced, less pipetting, and can lead to less tube-to-tube

variability. In short, using master mixes can lead to more reproducible results.

The following master mix recipe can be set up (using AccuPrime Pfx DNA

polymerase). This will differ depending on the polymerase used. Please also note

that you would need to scale up the amount of master mix depending on how

many reactions you are running. For example, if you are setting up 3 PCRs, you

would take the following recipe and multiply each component volume by 4 (3

reactions +1):

31

PCR Master mix (per 50 μl reaction)

Volume Component Final Concentration

34 μl Water

5 μl 10X AccuPrime Pfx

Reaction mix

1X

2 μl Forward Primer 100 pmol

2 μl Reverse Primer 100 pmol

1 μl MgCl2 or MgSO4 (50

mM)

1mM

1 μl AccuPrime Pfx

Polymerase (2.5U/ μl)

2.5U/50 μl

Note: The AccuPrime Pfx Reaction mix contains the reaction buffer AND

dNTPs at a final concentration of 300 μM.

The total volume of each master mix will be 45 μl to which 5 μl of template

DNA will be added before starting the PCR.

f. Amplify DNA using PCR (35 cycles for everything in bold):

PCR cycling conditions 94°C for 2 minutes Initial denaturation

94°C for 45 seconds Denaturation

55°C for 45 seconds Primer annealing

72°C for 90 minutes Primer extension

72°C for 5 minutes Final extension

g. Store on ice until ready for electrophoresis.

C. Separation of PCR product by electrophoresis

a. Using methods used before, perform gel electrophoresis to verify amplification by

PCR.

32

Mitochondrial DNA Analysis Using PCR

Introduction

As you recall, mitochondria are the energy-producing organelles of the cell and the

number of mitochondria per cell varies depending on the cell type, ranging from only a few in

skin cells to thousands in skeletal muscle cells. Unlike other organelles, mitochondria have two

distinct membranes. Porin proteins are present in the outer membrane, making it permeable to

ions and other molecules.By contrast, the inner membrane is enriched in a negatively charged

phospholipid known as cardiolipin, which helpsmake this membrane highly impermeable to

ions. Recall also that the inner membrane also contains the enzymes that constitute an electron

transport chain, the site of oxidative phosphorylation of adenosine diphosphate (ADP) to produce

adenosine triphosphate (ATP).

The DNA present in mitochondria is distinct from the DNA found in the cell’s nucleus.

Mitochondrial DNA (mtDNA) was the first part of the human genome to be sequenced. The

mitochondrial genome contains 16,569 base pairs of DNA that codes for 37 genes. MtDNA

encodes 13 polypeptides, all of which are subunits of the electron transport chain. However,

mtDNA does not encode the entire electron transport chain; for example, nuclear DNA encodes

for Complex II and subunits in the other complexes. Additionally, mtDNA codes for cytochrome

B (another constituent of the electron transport chain), and ATP synthase. One peculiarity of

mitochondrial protein synthesis is that mitochondrial mRNA uses a slightly different genetic

code than cytoplasmic translation. As such, mtDNA also encodes mitochondrial-specific

ribosomal RNA and transfer RNA. As all cells possess only one nucleus but several hundredor

thousand mitochondria, mtDNA is present in great excess over nuclear DNA in most cells. This

relative abundance of mtDNA is taken advantage of by forensic investigators after obtaining

crime scene specimens that are degraded or otherwise insufficient for nuclear DNA PCR

analysis. The D-loop region has a high degree of variability between individuals and can be

sequenced to demonstrate variations. MtDNA typing, however, cannot be used to conclusively

link suspects to crime scenes; rather, it can be used to include or exclude suspects from further

scrutiny.

During the past twenty years, an ever-increasingnumber of diseases have been shown to

be due to mitochondrial dysfunction. These disordersresult when mitochondrial ATP

generation isinsufficient to meet energy needs in a particulartissue. Because muscle and nerve

cells containlarge numbers of mitochondria, these organsystems are most affected by

mitochondrialdysfunction. Mitochondrial diseases may be due to mutations in mtDNA genes or

mutations in nuclear genes that encode mitochondrial enzymes. Diseases caused by mtDNA

mutations include the myopathies, diseases that affect various muscles, and

encephalomyopathies, which cause both muscular and neurological problems. Huntington’s

chorea, a devastating disease that results in dementia and loss of motor control, may be linked to

defects in oxidative phosphorylation caused by damage to mitochondria in neuronal tissues.

33

Other diseases such as Alzheimer’s and Parkinson’s disease involve mitochondrial

abnormalities, although it is unclear how these abnormalities relate to disease pathology.

Mitochondria also appear to play roles in aging and in apoptosis.

Since mitochondria are present in the cytoplasm, they are inherited independently from

the nucleus. A female egg cell possesses over 10,000 mitochondria, while a sperm cell has very

few. Thus, during fertilization, mitochondrial DNA is inherited almost exclusively from the

mother. Although a small amount of paternal mtDNA is present in the fertilized egg, this DNA

appears to be selectively destroyed by the newly fertilized egg. This pattern of inheritance of

mtDNA is known as maternal inheritance. Maternal inheritance is indicatedwhen all offspring,

male and female, ofthe mother are afflicted with a specific condition. The severity ofany

particular mitochondrial disorder ishighly variable, depending on the number of mutated

mitochondria inherited from the mother.

To examine mitochondrial DNA, the Polymerase Chain Reaction (PCR) is usually employed.

PCR was invented in 1983 by Dr. Kary Mullis at the Cetus Corporation in California. The

enormous utility of the PCR method is based on its ease of use and its ability to allow the

amplification of small DNA fragments.

Before performing PCR, template DNA is extracted from various biological sources.

Because PCR is very sensitive, only a few copies of the gene are required. Nevertheless, freshly

isolated DNA will provide better amplification results than older DNA specimens that may have

become degraded. In order to amplify the specific DNA or target sequence, two primers (short

& synthetic DNA molecules) are designed to correspond to the ends of the target sequence.

To perform PCR, the template DNA and a molar excess of primers are mixed with the

four “free” deoxynucleotides (dATP, dCTP, dGTP, and dTTP) and a thermostable DNA

polymerase. The most commonly used DNA polymerase is Taq DNA polymerase. This enzyme,

originally purified from Thermus aquaticus (a bacterium that inhabits hot springs), is stable at

very high temperatures. These components (template DNA, primers, the four deoxynucleotides,

and Taq DNA polymerase) are mixed with a buffer that contains Mg+2, an essential cofactor for

Taq polymerase. The PCR reaction mixture is subjected to sequential heating/cooling cycles at

three different temperatures in a thermal cycler.

In the first step, known as “denaturation”, the mixtureis heated to near boiling (94°C -

96°C) to “un-zip” (ormelt) the target DNA. The high temperature disrupts the hydrogen bonds

between the two complementary DNA strands and causes their separation. In the second step,

known as “annealing”, the reaction mixture is cooled to 45°C - 65°C, which allows the primers

to base pair with the target DNA sequence. In the third step, known as “extension”, the

temperature is raised to 72°C. This is the optimal temperature at which Taq polymerase can add

nucleotides to the hybridized primers to synthesize the new complementary strands. These three

steps - denaturation, annealing, and extension - constitute one PCR “cycle”. Each PCR cycle

doubles the amount of the target DNA in less than five minutes. In order to produce enough

DNA for analysis, twenty to forty cycles may be required. To simplify this process, a

34

specialized machine, called a thermal cycler, was created to rapidly heat and cool the samples.

In this experiment, students will examine their mtDNA from their own cells. To do this, PCR is

used to amplify two separate regions of the mitochondrial chromosome. Amplification of these

regions will result in PCR products of 921 and 672 base pairs. Following PCR, the amplified

DNA is analyzed using agarose gel electrophoresis.

Materials

• Thermo cycler

• Screw-capped tubes

• Microcentrifuge tube

• Ice Bucket

• Test tube rack

• Water bath

• PCR primers

• PCR Edvobead

o DNA polymerase

o Polymerase buffer

o dNTP mix

• Saline solution

Methods

D. Isolation of DNA from cheek cells

a. Label a 1.5 mL screw top microcentrifuge tube and a cup with your lab group

and/or initials.

b. Rinse your mouth vigorously for 60 seconds using 10 mL of the saline solution.

Expel the solution into cup.

c. Swirl the cup gently to resuspend the cells. Transfer 1.5 mL of solution into the

labeled tube.

d. Centrifuge the cell suspension for 2 minutes at full speed to pellet the cells. Pour

off the supernatant, without disturbing the cell pellet. Repeat steps b and c

twice more.

e. Resuspend the cheek cells in 140 μL of lysis buffer by pipetting up and down.

f. Cap the tube and place in a water bath float. Incubate the sample in a 55°C water

bath for 15 minutes.

g. Mix the sample by flicking the tube vigorously for 20 seconds.

h. Incubate the sample in a 99°C water bath for 15 min. Be sure to use screw-cap

tubes when boiling DNA isolation samples.

i. Centrifuge the cellular lysate for 2 minutes at low speed (6000 rpm).

j. Transfer 80 μL of the supernatant to a clean, labeled microcentrifuge tube.

35

k. Place the tube on ice or store the tube at -20°C if PCR is to be done at a later time.

E. Amplification of the mitochondrial regions

a. Label a 0.2 mL PCR tube with the sample and your initials.

b. Add 20 μL Mitochondrial primer mix, 5 μL extracted DNA (or control DNA) and

the PCR EdvoBead to the labeled 0.2 ml tube. At least one control reaction

should be performed per class to confirm that PCR was successful.

c. Mix the PCR sample. Make sure the PCR EdvoBead is completely dissolved.

d. Centrifuge the sample for a few seconds to collect the sample at the bottom of the

tube.

e. Alternatively, use the master mix protocol from the previous lab.

f. Amplify DNA using PCR. The conditions below in bold will run for 25 cycles.

PCR cycling conditions 94°C for 4 minutes Initial denaturation

94°C for 60 seconds Denaturation

55°C for 60 seconds Primer annealing

72°C for 2 minutes Primer extension

72°C for 5 minutes Final extension

g. After PCR, add 5 μL of 10x Gel Loading Solution to the sample.

h. Place tubes on ice or store at -20°C if electrophoresis is to be done at a later time.

F. Separation of PCR product by electrophoresis

a. Using methods used before, perform gel electrophoresis to verify amplification by

PCR.

36

Determination of Protein Molecular Weight Using SDS-PAGE

Introduction

As you recall, electrophoresis is a process in which molecules are exposed to an electric

field and separated on the basis of how they move through a substrate within the field. You have

already worked with agarose to separate DNA and in this lab, you will work with another type

of gel in order to separate proteins. Sodium dodecyl sulfate polyacrylamide gel electrophoresis

(SDS PAGE) allows the separation of proteins based on their molecular weight. This technique

can be used to determine whether a given protein is present in a sample, and/or to determine the

purity of the protein preparation, to estimate the approximate quantity of a protein, or even to

measure the size of a protein.

Polyacrylamide is used for proteins, particularly because it provides the right

environment where the protein(s) will not be denatured any more than they already need to be.

This allows different sized proteins to travel down the gel at different rates. Because different

protein molecules may be separated during the electrophoresis process, the goal is to start with a

linear structure. Therefore, the SDS is used to dissolve any hydrophobic molecules, which

results in a denatured, linear structure. Since proteins do not have a specific charge the way

DNA does, the SDS is also important because it contains a negative charge that is conferred onto

the protein (when it binds to it). This allows the protein to run “in the right direction” from

negative to positive. Once separated, the gel is stained and then several procedures can be done

including transferring the gel bands onto a membrane for detection by western blotting and/or cut

out and extracted for analysis by mass spectrometry.

The gels cast in SDS PAGE are casted much differently than in agarose electrophoresis.

The prepared acrylamide solution is poured into the thin space between two glass or plastic

plates that form a "cassette". Once the gel polymerizes, the cassette is vertically mounted into an

apparatus so that the top and bottom edges are placed in contact with buffer chambers containing

a cathode and an anode, respectively. The rest of the process is similar to agarose

electrophoresis in that the running buffer contains ions that conduct current through the gel.

When proteins are added in wells at the top edge and current is applied, the proteins are drawn

by the current through the matrix slab and separated by the sieving properties of the gel.

Typically, a “stacking” gel is cast over the top of the “resolving” gel in order to obtain optimal

resolution of proteins. The stacking gel has a lower concentration of acrylamide and a lower pH.

This allows the proteins in a loaded sample to be concentrated into one tight band during the first

few minutes of electrophoresis before entering the resolving portion of a gel.

Materials

• Pre-cast polyacrylamide gels

• Vertical gel electrophoresis box

• Power supply

• Hot plate

37

• DI water

Methods

A. Preparation of protein samples

a. Bring a beaker of water, covered with aluminum foil, to a boil. Remove from

heat.

b. Make sure the sample tubes A through D are tightly capped and well labeled. The

bottom of the tubes should be pushed through the foil and immersed in the boiling

water for 5 minutes. The tubes should be kept suspended by the foil.

c. Proceed to loading the gel while the samples are still warm.

d. Any unused protein samples should be frozen to be used for another experiment.

B. Preparation of precast polyacrylamide gels

Depending on the manufacturer of the precast gels, the following may differ

slightly:

a. Open the pouch containing the gel cassette with scissors. Remove the cassette and

place it on the bench top with the front facing up.

i. Note: The front plate is smaller (shorter) than the back plate.

ii. Some cassettes will have tape at the bottom of the front plate. Remove all

of the tape to expose the bottom of the gel to allow electrical contact.

iii. Insert the Gel Cassette into the electrophoresis chamber.

iv. Remove the comb by placing your thumbs on the ridges and pushing

(pressing) upwards, carefully and slowly.

b. Place the gel cassette in the electrophoresis unit in the proper orientation. The

protein samples will not separate in gels that are not oriented correctly.

c. Add the diluted buffer into the chamber. The sample wells and the back plate of

the gel cassette should be submerged under buffer.

d. Rinse each well by squirting electrophoresis buffer into the wells using a transfer

pipet.

e. The gel is now ready for practice gel loading and/or samples.

C. Preparation of protein samples

a. Place a fresh fine tip on the micropipet. Aspirate 20 μl of practice gel loading

solution.

b. Place the lower portion of the fine pipet tip between the two glass plates, below

the surface of the electrode buffer, directly over a sample well. The tip should be

at an angle pointed towards the well. The tip should be partially against the back

plate of the gel cassette but the tip opening should be over the sample well.

c. Do not try to jam the pipet tip in between the plates of the gel cassette.

38

d. Eject all the sample by steadily pressing down on the plunger of the automatic

pipet.

e. Do not release the plunger before all the sample is ejected. Premature release of

the plunger will cause buffer to mix with sample in the micropipet tip. Release the

pipet plunger after the sample has been delivered and the pipet tip is out of the

buffer.

f. Before loading protein samples for the actual experiment, the practice gel loading

solution must be removed from the sample wells. Do this by filling a transfer

pipet with buffer and squirting a stream into the sample wells. This will displace

the practice gel loading solution, which will be diluted into the buffer and will not

interfere with the experiment.

D. Loading protein samples

a. Change pipet tips between loading each sample. Make sure the wells are cleared

of all practice loading solution by gently squirting electrophoresis buffer into the

wells with a transfer pipet.

b. Two groups will share each gel. The pre-stained protein samples should be loaded

using the following lane assignments:

Group 1

Lane 1 20 µL of Tube A (Standard protein marker)

Lane 2 20 µL of Tube B (Unknown protein 1)

Lane 3 20 µL of Tube C (Unknown protein 2)

Lane 4 20 µL of Tube D (Unknown protein 3)

Group 2

Lane 1 20 µL of Tube A (Standard protein marker)

Lane 2 20 µL of Tube B (Unknown protein 1)

Lane 3 20 µL of Tube C (Unknown protein 2)

Lane 4 20 L of Tube D (Unknown protein 3)

E. Gel electrophoresis

a. After the samples are loaded, carefully snap the cover all the way down onto the

electrode terminals.

b. Insert the plug of the black wire into the black input of the power supply (negative

input). Insert the plug of the red wire into the red input of the power supply

(positive input).

c. Set the power supply at the required voltage and run the electrophoresis for 75

minutes at 125 volts. When the current is flowing, you should see bubbles

39

forming on the electrodes. Any sudsing that is seen is due to the SDS in the

buffer.

d. After the electrophoresis is finished, turn off power, unplug the unit, disconnect

the leads and remove the cover.

40

Bradford Protein Assay

Introduction

There are several ways to determine protein concentration. The Bradford assay is a rapid

and accurate way to measure protein concentration using spectrophotometric analysis.

Typically, in this assay, Coomassie Brilliant Blue is the dye used. It exists in three forms: a blue

anionic form, a green neutral form, and a red cationic. When protein binds to Coomassie, it

stabilizes the blue form. When Coomassie dye binds protein in an acidic medium, an immediate

shift in absorption maximum occurs from 465nm to 595nm with a color change from brown to

blue. Therefore, as protein concentration increases, blue color increases. Protein concentrations

are estimated by reference to absorbances obtained for a series of standard protein dilutions,

which are assayed alongside the unknown samples. The Thermo Scientific Coomassie Plus Kit

is a fast Coomassie-binding, colorimetric method for determining total protein concentration.

This method is a modification of the Bradford method and results in significantly less protein-to-

protein variation than is observed with other Bradford-type Coomassie formulations.

Materials

• Bovine Serum Albumin (BSA)

• Sterile water

• Test tube

• Spectrophotometer (Spec 20)

• Coomassie Plus Reagent

Methods

1. Pipette 0.05 mL of each standard or unknown sample into appropriately labeled test

tubes.

2. Add 1.5 mL of the Coomassie Plus Reagent to each tube and mix well.

3. For the most consistent results, incubate samples for 10 minutes at room temperature.

4. With the spectrophotometer set to 595 nm, zero the instrument on a cuvette filled only

with water. Then, measure the absorbance of all the samples.

5. Prepare a standard curve by plotting the measurement for each BSA standard vs. its

concentration in μg/ml. When preparing the standard curve, use a point-to-point as

opposed to a linear fit line. Use the standard curve to determine the protein concentration

of each unknown sample.

41

Vial Volume of

Diluent Volume and Source of BSA Final BSA

Concentration Final BSA

Concentration

A 0 300 µl of Stock 2000 µg/mL

B 125 µl 375 µl of Stock 1500 µg/mL

C 325 µl 325 µl of Stock 1000 µg/mL

D 175 µl 175 µl of vial B dilution 750 µg/mL

E 325 µl 325 µl of vial C dilution 500 µg/mL

F 325 µl 325 µl of vial E dilution 250 µg/mL

G 325 µl 325 µl of vial F dilution 125 µg/mL

H 400 µl 100 µl of vial G dilution 25 µg/mL

I 400 µl 0 0 µg/mL