Introduction To Molecular Biology

Life Science 3

Lab Component

Fall 2000

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TABLE OF CONTENTS Contents page Cover 1 Table of Contents 2 Laboratory Safety 3 Introduction 4 Laboratory Procedures 4 Pipetting 4 Pipetter Exercise 7 Lab 1: Experiment 1: Analysis of Proteins by SDS-PAGE 9 Instructions for constructing a standard curve in EXCEL 12 Lab Assignment #1 – Analysis of Proteins by SDS-PAGE 14 Lab 2: Experiment 2: Biochemical Assay of β-Galactosidase Activity 15 Lab Assignment #2 – Biochemical Assay of β-Galactosidase 22 Introduction to Labs 3 and 4 24 Lab 3: Experiment 1 (Part 1): Restriction Enzyme Analysis 28 Experiment 2 (Part 1): Identification of Bacteria Strains 29 Experiment 3 (Part 1): Growth of Bacteriophage 30 Lab Assignment #3 – Plasmid, Bacteria and Bacteriophage Week 1 32 Lab 4: Experiment 1 (Part 2): Restriction Enzyme Analysis 34 Experiment 2 (Part 2): Identification of Bacteria Strains 36 Experiment 3 (Part 2): Growth of Bacteriophage 36 Lab Assignment #4 – Plasmid, Bacteria and Bacteriophage Week 2 37 Appendix: Molecular Visualization with RASMOL 40

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LABORATORY SAFETY Safety in a molecular biology laboratory is of prime importance. One of the most important basic rules is NO FOOD AND DRINK IN THE LAB! You will be dealing with a variety of potentially hazardous chemicals as well as sophisticated instrumentation. The misuse of these chemicals or the instruments may result in harm to you or to your coworkers. Although it is necessary to be always cautious, this should not prevent you from enjoying the work and performing the experiments expeditiously. CLOTHING: A laboratory coat will serve to protect you and your clothing. Please provide your own laboratory coat. Although it is not compulsory to have a lab coat, many upper division lab courses require you to have one, so now may be a good time to buy one. FIRE: Fire extinguishers are located next to the doors in the laboratory. There are also extinguishers in the halls. If it is necessary to extinguish a flame, the first thing to remember is to stay calm. Many small fires may be extinguished by placing a non-flammable cover over the vessel so that oxygen cannot feed the fire. If you do use the extinguisher remember to remove the pin. EARTHQUAKE: Do not panic. Remain calm during minor earthquakes. For larger earthquakes, try and find cover under a table. Do NOT run out of the lab during an earthquake. After a large earthquake has stopped, turn off the gas from any Bunsen burners that are on and leave the lab by walking. Locate the nearest stairs and exit the building. Do not use the elevators. After a large earthquake assemble in the Life Sciences quadrangle. SPILLS AND OTHER ACCIDENTS: Always clean up any spills immediately, whether on your desk or any other place in the laboratory. If someone leaves a spill, it might contaminate your experiment, or cause an accident. If you are not sure of how to clean the spill, ask your teaching assistant. If you or anyone else in the laboratory is seriously cut, burned or hurt in some way, the teaching assistant or lab instructor should be informed. DO NOT DISCARD SHARP, POTENTIALLY DANGEROUS, OBJECTS IN THE TRASH. IF UNSURE, ASK YOUR T.A. BEFORE DISPOSING.

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INTRODUCTION LABORATORY PROCEDURES: This is a new course. A number of the problems have been ironed out, but there will no doubt still be problems and the laboratory course may not go as smoothly as planned, so we would appreciate your patience and understanding. PIPETTING These exercises will help you learn how to use a pipetter. All scientific labs around the world use the metric system. You should be familiar with the following prefixes: pico (p), nano (n), micro (µ) and milli (m) and the following standard units: Standard Units: 1000 pg = 1 ng 1000 pl = 1 nl 1000 ng = 1 µg 1000 nl = 1 µl 1000 µg = 1 mg 1000 µl = 1 ml 1000 mg = 1 g 1000 ml = 1 l Using the Pipetter: The pipetter is a volumetric instrument designed to measure and dispense liquids precisely and safely. The pipetter has a digital volumeter that displays the volume. The volume is adjusted by turning the black knurled adjustment ring (see Fig. 1B) and is continuously adjustable within the volume range for the pipetter. The maximum volume for the pipetter is shown on the pushbutton (see Fig. 1A). The pipetter uses disposable polypropylene tips (see Fig. 1E). The disposable tips ensure maximum safety for the user and no cross contamination between samples. To protect the user from contamination by the tips, the pipetter is equipped with a built-in tip ejector (see Fig. 1D). The volumeter displays three numbers and is read from top to bottom. The three numbers indicate the volume selected. The volume of the pipetter is set by turning the black knurled adjustment ring (see Fig. 1B). You will be assigned three pipetters. One dispenses volumes ranging from 100 – 1000 µl (p1000), the second from 20 - 200 µl (p200), and the third from 2 - 20 µl (p20). On a p1000, the number in red represents thousands of microliters. However, keep in mind that the maximum volume the instrument can handle, without causing damage, is 1000 µl or 1 ml (the red digit reads 1). For your purposes in lab, the red digit typically will be set to zero. The middle digit on the p1000 represents hundreds of microliters and the lowest digit tens. Thus, a volumeter setting of 010 (read from top to bottom) indicates a volume of 100 µl or the smallest volume that can be pipetted accurately. p1000 pipetters must be used with the large blue tips.

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On a p200, the top number indicates hundreds of microliters, the middle digit tens, and the lower digit individual microliters. The top digit should never be set greater than 2, since the maximum volume of a p200 is 200 µl (the volumeter reads 200 from top to bottom). The lower limit of the p200 is 20 µl, therefore, the lowest setting is 20 (a volumeter reading of 020). Finally, on the p20 pipetter, the top digit indicates tens of microliters, the middle individual microliters, and the red digit tenths of microliters. A volumeter setting of 200 indicates the maximum allowable volume of 20 µl, and a setting of 020 indicates the minimum volume of 2 µl. The p200 and p20 both use yellow tips.

Figure 1. The PIPETTER

Place a tip on the shaft of the PIPETTER. Press the tip on firmly using a slight twisting motion to ensure a positive, airtight seal. NEVER handle a liquid with a pipetter that has not been fitted with a tip.

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Press the pushbutton to the first positive stop (see Fig. 2A). Holding the pipetter vertically, immerse the tip into the sample liquid. Release the pushbutton slowly and smoothly to aspirate the sample (see Fig. 2B). Wait one second and then withdraw the tip from the liquid. Once the sample is aspirated, it is important to continue to hold the pipette vertically. Place the end of the tip against the inside wall of the tube/vessel at an angle of 10 - 40 degrees. Press the pushbutton smoothly to the first stop (see Fig. 2C). Wait one second. Press the pushbutton to the second stop to expel any remaining liquid (see Fig. 2D). Keeping the pushbutton pressed to the end, remove the pipetter by drawing the tip along the inside surface of the tube/vessel. Release the pushbutton (see Fig. 2E). Eject the tip by pressing the tip ejector button (see Fig. 2F). It is necessary to change the tip if a different concentration or type of liquid is being sampled next.

Figure 2. Operation of the PIPETTER

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Name: _______________________

Date: ________________________

TA: _________________________

Section: ______________________

PIPETTER EXERCISE: The ability to pipette well and accurately is central to the successful execution of LS 3 labs this quarter. Review the proceeding section called “Using the Pipetter” and Figures 1 & 2 before beginning these exercises. Table 1: Maximum and minimum allowable volumes for each pipetter. The pipetters will not be accurate at volumes outside of these ranges.

Pipetter Minimum volume (µl) Maximum volume (µl) P20 2 20

P200 20 200 P1000 100 1000

Exercise 1: Consider the following volumes: 350 µl

3.5 µl 35 µl Which pipetter should be used to measure the above volumes?

Indicate the setting in each volumeter window. How do your pictures compare?

3.5 µl 35 µl 350 µl

Top Window Bottom Window

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Exercise 2: Obtain 3 microcentrifuge tubes from your TA and using the colored water provided on your tray:

i. Using the appropriate pipetter, aspirate 3.5 µl of colored water and transfer to a microcentrifuge tube.

ii. Trade microcentrifuge tubes with others in your group. iii. Aspirate 3.5 µl of colored water from one of your lab partners’ microcentrifuge tubes.

Repeat the steps above using a 35 µl volume and a 350 µl volume. Answer the following questions for all three volumes, 3.5, 35 & 350 µl: Is there any liquid left over in the microcentrifuge tube? Did you run out of sample to aspirate from the microcentrifuge tube? What are some possible reasons why you may have answered yes to the

above questions?

Exercise 3: Pipette check out. Demonstrate your pipetting skills to your UA or TA and earn their signature of approval.

__________________________ Signature

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LAB 1 – ANALYSIS OF PROTEIN SIZE AND SUBUNIT COMPOSITION USING SDS-POLYACRYLAMIDE GEL

ELECTROPHORESIS

Introduction: Some proteins contain more than one polypeptide chain (subunit) arranged in quarternary structure. There are many different types of such multimeric proteins, for example dimers, tetramers, pentamers and hexamers. These may be composed of identical polypeptide chains, as in a homodimer, or two (or more) different polypeptide chains as in a heterodimer. The forces that stabilize the interaction between subunits are similar to those that stabilize tertiary structure of a single-subunit protein; they contain closely packed nonpolar sidechains, hydrogen bonds and in some cases intermolecular disulfide bonds. There are several methods for estimating the size of proteins. Two methods, gel filtration and SDS-Polyacrylamide gel electrophoresis (SDS-PAGE), yield different but complimentary data. Gel filtration provides an estimate of the molecular weight of the protein in its ‘native’ state, i.e. its intact and function form. In contrast, the detergent SDS denatures proteins, disrupting noncovalent linkages between subunits (polypeptides). When SDS is used with a reducing agent that breaks disulfide linkages, the individual subunits of the protein can be separated, on the basis of differences in their molecular weights, using polyacrylamide gel electrophoresis (PAGE). The summed weight of the individually isolated subunits should equal the molecular weight of the protein as estimated using gel filtration. Discrepancies in the molecular weight estimates from these two techniques provide important information about the subunit composition of a protein. For example, if the molecular weight of a protein is determined via gel filtration to be 600 daltons, but the weight of the protein as determined by isolating individual subunits using SDS-PAGE indicates a total weight of 200 daltons, the most parsimonious explanation for this discrepancy is that the protein consists of three identical subunits each weighing 200 daltons (3 x 200 daltons = 600 daltons). How does the SDS-PAGE technique actually work? Electrophoresis is a method for separating charged molecules, such as proteins and nucleic acids, in an electric field. The rate of a molecule’s migration through an electrical field depends on the net charge, the size and shape of the molecules and the strength of the electric field. As an analytical tool electrophoresis is used to estimate the size of molecules. SDS-PAGE, the most common type of electrophoresis applied to proteins, is also referred to as denaturing electrophoresis. The molecular weight is estimated by comparing the migration of a protein in its fully denatured or unfolded state to that of other standard proteins that are also fully denatured. “SDS” refers to the anionic detergent sodium dodecyl sulfate (also called sodium lauryl sulfate; use of the latter, Latin form of the name is common in the shampoo industry). The protein of interest is mixed with a loading buffer that contains a reducing agent and SDS. The reducing agent breaks disulfide linkages. The SDS disrupts the noncovalent interactions

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between subunits of proteins, causes each polypeptide chain to unfold, and the SDS then associates with the unfolded backbone of the protein, conferring an overall negative charge to the protein in proportion to its length. The average amount of SDS associated with a protein is estimated to be one molecule of SDS/two amino acid residues. Thus, proteins in sample buffer effectively become rods of negative charge with equal amounts of charge per unit length. “Polyacrylamide” in PAGE refers to the gel matrix in which the proteins migrate. The gel is mounted between two buffer chambers containing separate electrodes so that the only electrical connection between the two chambers is through the gel. The proteins are loaded into “wells” at the end of the gel with the negative electrode. Standard proteins of known size are loaded into at least one well. Current is then applied and the proteins migrate toward the positive electrode at a rate proportional to their size. The electrophoresis run is completed when the smallest, and thus, fastest migrating proteins reach the bottom of the gel. The proteins can be visualized in several different ways. The most typical is by immersing the gel in a stain, Coomassie blue, which binds to the proteins. In this lab, all proteins have been covalently linked to a fluorescent tag—fluorescein isothiocyanate or FITC. FITC is a small (389 dalton) molecule that can be attached via its isothiocyanate group to the amino terminal and primary amines in proteins, especially lysine. This tag, and therefore the protein bands on the gel, become visible when illuminated with UV light. The sizes of the unknown protein subunits are determined by comparing the distances these polypeptides migrated to a standard curve constructed from the distances migrated by the standard proteins (i.e. proteins of known weight). The objective of this lab: In this lab, you will be given a protein of known molecular weight (predetermined by gel filtration on the native molecule) but unknown subunit composition. You will use the SDS-PAGE technique, in conjunction with the results of the gel filtration analysis (provided), to determine the size and number of the individual subunits of this protein. Grading: At the end of lab, each individual will turn in a sheet showing their standard curve and molecular weight estimation of their unknown protein. The report will also include answers to specific questions about the subunit composition of the unknown protein. RASMOL During the electrophoresis, you will learn to use the 3-D visualization program, RASMOL, to examine the 3-D structures of β-galactosidase, a tetrameric enzyme you will investigate in lab 2. You will also examine a single monomer of this protein. See the Appendix (p.40).

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Experimental Procedure 1. Obtain protein samples in loading buffer from your TA. Be sure to keep them on ice

until you are ready to use them. 2. Place samples in a 65°C bath for 5 minutes to fully denature the proteins. 3. Place samples on ice and immediately load 10µl into each well. Record which well

contains your sample. 4. Run gel at 130 V for 1 hour. 5. i. After your TA has separated the gel from the gel plates, take a photograph of the

gel using the gel documentation system. Place the gel in the center of the hood on the transilluminator. Turn on the power strip to engage the documentation system. After turning on the UV source in the transilluminator (bottom switch on the front), you should be able to view your gel on the monitor. If you encounter problems, consult your TA.

ii. Using the print button on the integration control unit, print a copy of the gel. This print can be cut to share with others in your group. iii. After you have made successful prints of your gel, turn off the UV light in the transilluminator as well as the power strip for the documentation system. Discard the gel as directed by your TA. Clean the transilluminator with a paper towel.

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How to Construct a Standard Curve in EXCEL

On the photograph of the gel:

Measure the distances polypeptides (standards) in the marker lane ran. To do this take a ruler and measure the distance (in cm or mm) from the well to each of the bands in the lane.

Using EXCEL to construct and fit a curve:

1. Enter the distance each polypeptide standard migrated (x variable) and its mass (y variable) in two adjacent columns on the spread sheet.

2. Highlight both columns and click on graph wizard icon (upper row of

tool bar, looks like a vertical bar chart).

3. Select the X-Y (Scatter) plot from left column of plot types.

4. Select the chart sub-type without lines (Scatter plot without lines - top box).

5. Follow directions to create the plot. Make sure you label your axes.

6. After points are plotted, select the graph by clicking on it.

7. Go to “Chart” on tool bar and select “Add trendline” from the drop-

down menu.

8. Select the “Logarithmic plot” box.

9. Move to “options” tab. Click “display equation on chart”.

10. Click “OK”.

11. Examine fit of curve to points and then try another curve.

12. Return to “Add trendline” function and try the power curve.

13. Compare the power and logarithmic curves on plot. Use the curve that mostly closely conforms to the distribution of points on the plot (fits the points best).

14. Delete the plot with the poorer fit. To do this, click on the curve to

select it and then hit delete on the keyboard.

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15. When you are satisfied with your fit, add your name to your figure in the title box and prepare to print your curve.

16. Click on the figure to make sure you have selected it. Go to “File” on

the tool bar and select print preview.

17. If you are satisfied with the preview, select print and ask your TA or UA to retrieve your figure from the printer.

Determine Size of Unknown Polypeptide(s):

1. Measure the distance that your unknown polypeptide(s) migrated. Again measure from the well to the band or to each band in the lane in which your unknown was run. Measure in the same units used to construct the standard curve.

2. Use the equation of your standard curve to determine the size of your

unknown polypeptide. Remember the x variable in the equation is the distance the unknown polypeptide(s) migrated and the y variable is what you want to know and what you solve the equation for – the size of the polypeptide.

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Name: ________________________ Date: _________________________ TA: __________________________ Section: _______________________ DUE: AT END OF LAB LAB ASSIGNMENT # 1 – Analysis of Proteins by SDS-PAGE SAMPLE ID#_________ Estimated Molecular Weight by Gel Filtration__________ Estimation of Molecular Weight by SDS-PAGE: 1. Paste the photo of your gel here. Use EXCEL to construct a standard curve. Then

determine the size of each of the polypeptides (subunits) on your gel. Please attach your standard curve.

Analysis of Subunit Composition of Sample Protein 2. How many subunits does your unknown protein contain? Indicate the number of

each subunit in the protein and the molecular weight of each of these subunits. 3. How could you use electrophoresis to determine if the subunits are associated by

disulfide bonds?

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LAB 2 - BIOCHEMICAL ASSAY OF β-GALACTOSIDASE ACTIVITY

Introduction Many types of enzymes, that perform all sorts of functions, are found in cells. Some enzymes are needed all the time (e.g., hexokinase), whereas others are needed only under certain conditions. For example, the enzyme β-galactosidase catalyzes the hydrolysis of lactose (a disaccharide found in milk) to galactose and glucose. However, bacteria growing in glucose (no lactose) have no need for the enzyme β-galactosidase. Is this enzyme still synthesized under these conditions? If so, how much? How can you tell? Enzyme activity is measured using a procedure called a biochemical assay. An assay is a method to indirectly assess how much of a given protein (enzyme) is produced per cell, per unit time. The ability to measure enzyme activity, like the ability to determine protein structure (introduced in Lab 1), is a technique critical to the investigation of the relationship between genes, proteins, and cellular function. How do you biochemically assay the activity of β-galactosidase? To measure the amount of enzyme in a cell, one needs to determine the amount of that product produced in a given time by a given number of cells. Since it is not easy to assay β-galactosidase activity by measuring the amount of galactose or glucose produced or the amount of lactose consumed, a derivative of lactose, o-NO2-phenyl-β-D-galactopyranoside (ONPG), is used as the substrate. ONPG is a colorless compound. However, cleavage of ONPG by β-galactosidase produces a yellow compound, o-NO2-phenol. This makes ONPG a useful chromogenic substrate for assaying β-galactosidase activity. The amount of o-NO2-phenol produced is measured by determining the absorption (optical density, OD) of a sample at 420 nm in a spectrophotometer.

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The amount in o-NO2-phenol produced per minute per bacterium reflects the amount of enzyme in a typical cell and is referred to as units of activity. For our purposes, units of enzyme activity will be defined as:

Units = OD420/ (time x volume x OD600) OD420 is the optical density (absorbance) of a sample at 420 nanometers and is proportional to the amount of o-NO2-phenol produced. Time is the length of time that the reaction of β-galactosidase and ONPG is permitted to proceed; it is measured in minutes. Volume is the volume of the sample (removed from a culture) that is assayed for β-galactosidase activity; it is measured in milliliters. OD600 is the optical density of a sample at 600 nanometers; it provides an estimate of cell density in the sample (i.e. the number of cells/ml). (An OD600 equal to 1 represents approximately 2 x 108 cells/ml.) The objective of this lab: In this experiment you will use the chromogenic substrate ONPG to assay β-galactosidase activity in E. coli cells grown in the presence of glucose and lactose, as well as in the absence of a sugar substrate (the control). The assay will allow you to determine (1) if E. coli synthesizes the enzyme β-galactosidase in the absence of lactose, the substrate upon which β-galactosidase acts and (2) if the induction of β-galactosidase synthesis in E. coli is time dependent. To accomplish this you will assay β-galactosidase activity at two time points during the experiment: (1) 15 minutes after E. coli cells are introduced to sugar (glucose or lactose) and (2) 90 minutes after the introduction. A few notes about the protocol: • Luria broth provides essential nutrients for bacterial growth. • Chloroform is added to partially disrupt the cell membrane, allowing the substrate

(ONPG) to diffuse into the cell. • Z buffer promotes the reaction between β-galactosidase and ONPG by optomizing the

pH of the sample. • Addition of Na2CO3 stops the reaction of β-galactosidase and ONPG by changing the

pH from 7.0 to 11. β-galactosidase is very pH sensitive. Grading: At the end of lab, each individual will turn in a sheet showing their measurements of cell culture growth and β-galactosidase activity. The report will also include answers to specific questions regarding your results.

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EXPERIMENTAL PROCEDURE 1. Obtain the following test tubes from your TA:

TEST TUBE A: 4.0 ml Luria Broth (LB). TEST TUBE B: 3.6 ml LB + 400 µl 4% glucose TEST TUBE C: 3.6 ml LB + 400 µl 4% lactose Note: The capital letters A, B, and C refer to these tubes throughout the remainder of the protocol. Lower case versions of these letters are used to identify subsamples of these original cultures.

2. TIME ZERO: ESTABLISH CULTURES AND β-GALACTOSIDASE BLANK

a. Set-up β-galactosidase Blank: i. Retrieve one microcentrifuge tube. Label this tube “β-gal blank”.

ii. Into this microcentrifuge tube, pipet 100 µl of Luria Broth from test tube A.

iii. Set aside until Step 3bii.

b. Establish E. coli cultures in test tubes A, B and C.

i. Retrieve E. coli from your TA.

ii. Pipet 400 µl of E. coli bacteria into each A, B, and C tube.

iii. Gently mix the E. coli cells and broth by closing cap tightly and inverting each tube (A, B, and C) 1-2 times.

iv. Label each tube (A, B, and C) with your group’s name.

v. Place them in the 37°C incubator. Note the time.

vi. Incubate the tubes for 15 minutes.

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3. TIME 15 MINUTES: BEGIN ANALYSES (See flow chart)

a. Subsample cultures A, B and C to determine E. coli cell density:

i. Retrieve 3 cuvets from your TA. Without writing on cuvets, label one cuvet a, one b, and one c.

ii. Retrieve tubes A, B and C from the incubator. Gently mix the E. coli cells and

broth in by closing cap tightly and inverting each tube (A, B, and C) 1-2 times.

iii. Remove 1 ml from tube A and place it in the cuvet a. Repeat for tubes B and

C. iv. Proceed to Step 4: Determine E. coli Cell Density

v. Note: While part of your group measures cell density, part of your group can

perform the ß-galactosidase assay (see step 3b immediately below).

b. Subsample cultures A, B and C to assay cultures for β-galactosidase activity:

i. Retrieve 3 microcentrifuge tubes from your TA. Label one tube “a”, one “b”, and one “c”.

ii. To each of these three microcentrifuge tubes (a, b, and c) AND to the “β-gal

blank” (see Step 2a), add 600 µl of Z buffer mix.

iii. Remove 100 µl from tube A and add it to microcentrifuge tube “a” containing Z buffer mix. Repeat for tubes B and C. Do not add anything to the β-gal blank.

iv. Proceed to Step 5: B-galactosidase Assay

c. Return tubes A, B, and C to the incubator.

4. DETERMINE E. COLI CELL DENSITY: a. Follow directions posted on the spectrophotometer (also see last page of lab). b. Be sure that spectrophotometer is set to read at 600 nm. c. Use the LB blank provided (not the “β-gal blank” containing Z buffer mix).

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d. Measure the absorbance (optical density, OD) of the 1 ml subsamples labeled a, b, and c that you placed in the cuvets in Step 3a.

e. Record your data (the values in the column headed 600λ). f. Discard sub-sample cuvets into bleach cup.

5. β-GALACTOSIDASE ASSAY:

a. To the 3 microcentrifuge tubes labeled a, b, and c (which contain 700 µl of E. coli - Z buffer mix) and to the “β-gal blank” (see Part 3bii), add 2 drops of chloroform. Chloroform should not be inhaled, therefore, this step should be done in the hood. Close caps tightly.

b. Vortex all 4 microcentrifuge tubes for 30 seconds.

c. After vortexing, let all 4 tubes sit 1-2 minutes, during this time the chloroform will

settle to the bottom of the tubes.

d. Transfer 650 µl of sample a into a clean, labeled microcentrifuge tube. Repeat for samples b and c and the “β-gal blank”. Remove only the top layer of liquid. Be careful not to remove any of the buffy layer of cell debris at the liquid interface or the chloroform below the interface at the bottom of the tube.

e. Place tube with remaining chloroform in the rack in the hood.

f. Add 140 µl of ONPG (4 mg/ml) to each of the 3 samples and the “β-gal blank”.

Mix by inverting each tube several times.

g. Incubate tubes at room temperature for 15 minutes. During this time β-galactosidase, if present, will cleave ONPG to produce the yellow product o-nitrophenol.

h. Add 350 µl of 1 M Na2CO3 to each of the 3 samples and the “β-gal blank”. Mix by inverting each tube several times. (Yellow color is stable after the addition of Na2CO3.) Na2CO3 stops the cleavage reaction.

i. Transfer 1 ml of sample a into a cuvet. Repeat for samples b and c and the “β-gal

blank”. Keep track of the sample identity in each cuvet but do not write on cuvets.

j. Measure the absorbance (optical density, OD) of a, b and c at 420 nm visible light using the “β-gal blank” as your blank. Record your data (the values in the column headed 420λ).

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k. Save the β-gal blank.

l. Discard sample cuvets a, b, and c into bleach cup with lids open. 6. TIME 90:

Repeat the steps above, starting at Step 3 (Time 15), to measure cell growth and β-galactosidase activity 90 minutes after time zero.

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CORRECT FLOW CHART FIGURE IS IN FILE: B-GAL FLOW CHART 3.

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BECKMAN DU 640B SPECTROPHOTOMETER OPERATING INSTRUCTIONS

FOR FIRST SAMPLE (BLANK): Turn on the monitor (Spectrophotometer should be on already). Under “Routine Measure” click “Fixed Wavelength” Next to “Sample ID” there is “λ” with a number following. Click on the first number and a keypad will appear. Change the number to the wavelength you want to measure. When you are finished, click “OK” Click “[VIS OFF]” so display is “[VIS ON]” (bottom, left of screen). Insert BLANK cuvet, close cover. Click “BLANK” (bottom, left of screen). When it is finished reading, remove BLANK cuvet. FOR ALL OTHER SAMPLES:

Insert SAMPLE cuvet, close cover. Click “Read Samples” (top, left on screen). Record Absorbance (Abs) numbers displayed for your chosen wavelength.

Insert next sample and follow the preceding steps. If there are no more samples to be read at this time, click “[VIS ON]” so display is “[VIS OFF]”. TO SWITCH BETWEEN WAVELENGTHS: Note: When you switch between wavelengths you must use a new blank.

After all data has been recorded from the PREVIOUS wavelength, click “SaveClear” (top, right on screen).

Click the box “Save results before clearing?” so that the box is clear. Click “OK” Next to “Sample ID” there is“λ” with a number following. Click on the first number and a keypad will appear. Change the number to the wavelength you want to measure. When you are finished, click “OK” Insert BLANK cuvet, close cover. Click “BLANK” (bottom, left of screen). When it is finished reading, remove BLANK cuvet. WHEN ALL SAMPLES ARE COMPLETED (AT THE END OF LAB): Remove last sample Click “[VIS ON]” so display is “[VIS OFF]”. Click “Quit” Click the box with a dot at “Save Results” so box is clear. Click “OK” Turn off the monitor.

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Name: ________________________

Date: _________________________ TA: __________________________ Section: _______________________ DUE: AT END OF LAB LAB ASSIGNMENT # 2 - Biochemical Assay of β-Galactosidase

A. Culture grown in Luria Broth (This is the control.) Time point OD600 Volume Time OD420 Units 15 min

90 min

B. Culture grown in Luria Broth + glucose Time point OD600 Volume Time OD420 Units 15 min

90 min

C. Culture grown in Luria Broth + lactose Time point OD600 Volume Time OD420 Units 15 min

90 min

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1. Is the level of β-galactosidase activity influenced by the carbon (sugar) source in the

media? Explain using your results. 2. Did you observe a difference in β-galactosidase activity between your 15-minute time

point and your 90-minute time point in all three cultures (A, B and C)? Explain your results.

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Introduction to Labs 3 and 4 Introduction Molecular Biology techniques rely heavily on analysis and manipulation of specific DNA segments. In order to obtain enough DNA to study, sequence, use as probes, etc., one must isolate relatively large quantities of the specific DNA segment. Amplification can be done in vivo by cloning the DNA into a vector (e.g., plasmid or phage) and amplifying it in bacteria, or in vitro using PCR (Polymerase Chain Reaction). We will focus here on cloning DNA fragments. DNA cloning became possible with the discovery of two types of enzymes: restriction enzymes and DNA ligases. Restriction enzymes cut DNA at specific short sequences generating a reproducible set of fragments. DNA ligases catalyze the formation of bonds between fragments of DNA, allowing the insertion of DNA into self-replicating DNA molecules or vectors. Together, these enzymes allow the creation of recombinant DNA molecules that can be introduced into cells (most often bacterial cells) and grown in large quantities. All the descendents from a single such cell will carry the same recombinant DNA molecule (a clone). This cloned DNA can then be isolated and manipulated in many different ways. Plasmids One of the most commonly used vectors is the bacterial plasmid. Plasmids are naturally occurring, small, circular, double stranded, extrachromosomal DNA molecules capable of autonomous replication. Plasmids that are used as vectors have been engineered to contain several essential items: a replication origin, a drug-resistance gene, and a region in which foreign DNA fragments can be inserted (a region containing restriction sites). pUC19, the plasmid you will be investigating in lab, contains an ampicillin resistance gene (ampr). Additionally, the insertion region lies within the lacZ gene that codes for β-galactosidase (Fig. 1). As you will see in lab, this is a useful feature because it allows one to use a simple biochemical assay to distinguish bacteria containing plasmids with inserts (pieces of foreign DNA to be cloned) from those without, a process called screening. In particular, bacteria containing plasmids with inserts will not synthesize β-galactosidase because the DNA fragment of interest has been inserted into the middle of the lacZ gene. This is detected by plating the bacteria on a glucose medium containing an indicator, the lactose derivative X-gal (5-bromo-4-chloro-3-indolyl-β− D-galactoside), along with IPTG (isopropylthio-β− D-galactoside, a molecule that induces E. coli to transcribe the lacZ gene but is not digested by β-galactosidase; thus IPTG continuously induces β-galactosidase synthesis in culture). The cleavage of X-gal by β-galactosidase produces a blue compound. Thus, colonies of bacteria derived from a successfully transformed bacterium will appear white. In contrast, bacterial colonies lacking the DNA insert will appear blue.

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Foreign DNA is inserted into a plasmid using two types of enzymes: restriction and ligation. Restriction enzymes (also called endonucleases) cleave double stranded DNA molecules at specific, short nucleotide sequences (restriction sites) that are unique to each enzyme. Some enzymes, for example EcoRI and HindIII, make staggered cuts in the DNA, producing “sticky ends” which are single stranded tails that lack complimentary base pairs and thus, will pair up with the “sticky ends” of other fragments generated using the same enzyme. Other restriction enzymes produce “blunt ends” in which all of the nucleotides at fragment ends are base-paired to nucleotides in the complimentary strand. Often plasmid vectors contain a polylinker, an insertion region (sequence) which contains one copy of each of several different restriction sites. This allows one to clone a variety of restriction fragments (e.g., fragments with EcoRI ends, HindIII ends, or fragments with one EcoRI end and one HindIII end) and to clone a fragment in a known orientation (e.g., from EcoRI to HindIII). Additionally, during a cloning experiment, digestion with two different restriction enzymes inhibits ligation of the vector without an insert because the two enzymes generate free ends with incompatible nucleotide sequences. DNA ligase is used to covalently bond the ends of restriction fragments. Thus, when inserting a piece of foreign DNA into a plasmid, both the plasmid and insert must be digested with the same restriction enzyme(s) to create complimentary fragment ends. These complimentary pieces of DNA then are bonded covalently using DNA ligase. Bacteria are considered transformed (or to have undergone transformation) when they successfully have acquired and are able to replicate the DNA vector. Transformation efficiency using plasmids is low; only a small fraction of bacteria successfully take up a plasmid. Consequently, it is necessary to distinguish transformed bacteria from those lacking the plasmid. This process, called selection, is made possible by the presence of the drug-resistant gene in the plasmid (e.g., ampicillin resistance in pUC19). Successfully transformed bacteria will grow on a medium containing the drug while those lacking the plasmid will not (i.e. bacteria with the plasmid are selected for while those lacking the plasmid are selected against under these particular conditions). Selection

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assays also include a control; bacteria are grown on a plate lacking the selective agent to ensure that the bacteria are viable. Collectively, selection and screening (described previously) allow one to determine: 1) which bacterial colonies have the plasmid (selection) and 2) of those colonies with the plasmid, which have plasmids containing the insert (screening) and, thus, have cloned the DNA of interest. Cloned DNA fragments are retrieved by digesting plasmids with the same restriction enzyme(s) that was (were) used during the original insertion. The relatively large pieces of plasmid DNA and the smaller, cloned fragments are then separated by electrophoresis on an agarose gel, a process analogous to the separation of different size polypeptides using polyacrylamide gel electrophoresis. As in PAGE, the fragments separate according to their relative sizes (however, in contrast to SDS-PAGE, size here refers to length in bases) and the smallest fragments migrate the farthest. The sizes of the cloned fragments are determined by comparing the distances these fragments migrated to a standard curve constructed from the distances migrated by “standards” or fragments of known length that are run on the gel simultaneously. The DNA is visualized by first incubating the gel in a solution of ethidium bromide. This highly mutagenic substance binds to DNA by intercalating between the stacked bases of DNA, producing highly concentrated patches of the dye in the exact location of DNA bands on the gel. When illuminated with UV light, the DNA bands exhibit much greater fluorescence relative to the residual dye in free solution. (Ethidium bromide absorbs UV-irradiation in 300 nm and 360 nm range and subsequently re-radiates this energy at 590 nm in the red-orange region of the visible spectrum.) In addition to facilitating DNA amplification, the ability to engineer plasmids and transform bacterial cells provides opportunities for other kinds of productive manipulations. For example, this technique is also used to produce large quantities of protein that is normally expressed at low rates as well as to isolate fragments of a particular nucleotide sequence from a mixture of many different sequences.

Bacteriophage The bacteriophage (phage) is the second type of vector that will be introduced in this lab. Phage are naturally occurring bacterial viruses that incorporate their DNA into that of a bacterium and co-opt the cell’s machinery to reproduce this DNA. Phage are useful alternatives to plasmids because 1) the infectious nature of phage greatly increases the efficiency of bacterial transformation making it possible to clone larger numbers of DNA fragments rapidly and 2) phage vectors can incorporate slightly larger DNA fragments than plasmids (25 versus 20 kb respectively). Phage lambda (a commonly used vector) has two growth pathways, lysogenic and lytic. In the lysogenic pathway, viral DNA is incorporated into the host cell genome and is replicated only when the host cell divides producing a single copy in each daughter cell. In the lytic pathway, many new virus particles (virions) are synthesized without bacterial

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cell division. These particles accumulate in the cell and eventually cause it to rupture (lysis), releasing up to 100 virions into the surrounding environment. In phage that have been engineered to be vectors, the genes coding for the lysogenic pathway are replaced with the DNA fragment of interest. Thus, infected bacterial cells must enter the lytic pathway, eventually rupturing and releasing new virions to infect surrounding cells. When a phage-infected bacterial culture is plated out in a petri dish, distinct holes, approximately 2 mm in size, appear in the resulting lawn of bacteria. These holes, called plaques, contain dead bacterial cells that were infected and lysed by phage. Each plaque is started when one phage infects one bacterium. The released progeny infect neighboring cells, which subsequently release more phage. The final result is about 1 million phage per plaque, each of which contains a copy of the DNA fragment of interest. Phage are used often to produce DNA libraries (a set of clones that contains every sequence in the genome of a particular organism). In this process, the genome is partially digested using a restriction enzyme and the fragments are ligated into phage. The end result is that each phage contains a different fragment of the genome (fragments may contain portions of sequences found in other fragments but each fragment is virtually an unique piece of DNA). Incubation of these phage with bacterial cells, for a time shorter than one lytic cycle, ensures that each subsequent plaque contains multiple copies of a distinct DNA fragment (i.e. the DNA in each plaque differs from that in all other plaques). In contrast, multiple plaques may contain the same DNA sequence if bacteria are incubated with phage for longer than one lytic cycle because cell lysis introduces many copies of a single sequence into the incubation.

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LAB 3 Introduction: Labs 3 and 4 are contiguous. You will initiate three separate experiments (1, 2 and 3) during Lab 3 and you will conclude these experiments during the following lab period (Lab 4). A separate, graded assignment is due at the end of each lab period. The objectives of labs 3 and 4:

1. To use a restriction digest and gel electrophoresis to distinguish two types of plasmids:

a. a plasmid b. a plasmid plus insert

and to estimate the length of the insert.

2. To select for and screen bacteria in order to distinguish between bacterial strains with :

a. no plasmid b. plasmid c. plasmid plus insert

3. Compare growth of a phage plaque to a bacterial colony.

Grading Lab 3: At the end of lab, each individual will turn in the assignment stating their choice of media for differentiating between bacterial strains and the expected results of the selection/screening experiment. The assignment also will include answers to specific questions regarding plasmid and phage vectors.

EXPERIMENT 1 (Part 1) - Restriction Enzyme Analysis of Plasmids Each group will be given two tubes of DNA. Each contains the plasmid vector pUC19 (see Figure 1). However, in one tube, pUC19 has a DNA insert cloned between the EcoRI site and HindIII site. In the other, pUC19 does not have a DNA insert. Your goal is to determine which of the two tubes contains the plasmids with the insert and to estimate the length of the insert. Experimental Procedure 1. Using one of the labels provided, write the section, group and digestion letter and place

on the lid of the restriction digest tube. Also, with a marker, write directly on the side of the microcentrifuge tube in case the lid label comes off.

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2. To each tube (that already contains 2 µl plasmid DNA) add:

• 14 µl H20 • 2 µl 10x restriction buffer • 1 µl EcoRI (5 units) • 1 µl HindIII (5 units)

3. Mix the components of this restriction digest by tapping the microcentrifuge tube a

few times. 4. Briefly spin the samples for 30 seconds using a microcentrifuge. (Be sure to balance

the centrifuge before spinning.) 5. Place both restriction digests in the 37°C incubator. Incubate for 45 minutes 6. After 45 minutes, place your digest tubes in the freezer. You will separate the

restriction fragments by electrophoresis during Part 2 of this experiment, which will be conducted during the next lab period (Lab 4).

EXPERIMENT 2 (Part 1) - Identification of Bacteria Strains Using Selection and Screening

Each group will be given three strains of E. coli (from a standard JM101 culture) labeled Α, Β and C. You will determine which strain (A, B, and C) corresponds to which of the following:

1. Strain that does not contain the plasmid vector pUC19. 2. Strain that contains pUC19 but no insert. 3. Strain that contains pUC19 with an insert.

Experimental Procedure 1. After the class discussion about E. coli plasmids and strains, each group should decide

what type of plate upon which to streak the three bacterial strains. Your choices are:

1. Luria Broth (LB) media (rich media that does not contain lactose) 2. LB media + ampicillin 3. LB media + ampicillin + XG 4. LB media + ampicillin + XG + IPTG

2. Confirm your decision about which plates to use with your TA or UA before retrieving

plates.

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3. Review the computer video on streaking before beginning to streak your plates. Good

sterile technique is always critical when working with bacteria. 4. Use a sharpie marker to draw a “Y” on the BOTTOM (agar containing side) of the

plate to designate the sectors onto which you will streak each strain (A, B and C). Label each sector with a letter (A, B or C).

5. Use the sterile loop to gently streak the cells in a zigzag path across the surface of the

appropriate sector. Be careful not to gouge the agar. The aim is to streak out the bacteria so that you are able to isolate single colonies.

6. Turn the plates upside down and label the plates around the bottom edge with the

following information: date, section number and group. 7. Your TA will put the plates for the entire class in the 37°C air incubator UPSIDE

DOWN (lid on bottom; this prevents moisture dripping onto the agar). The plates will be incubated overnight and then stored in the refrigerator until the following lab (Lab 4) when you will complete Part 2 of this experiment.

EXPERIMENT 3 (Part 1) - Growth of Bacteriophage In order to gain a first-hand knowledge of the difference between a clone of phage (also known as a plaque) and a clone of bacteria (also known as a colony) you will inoculate bacteria with the phage λ (lambda) and observe the resulting plaques which form on an agar plate. As in the previous experiment, sterile techniques are very important. Experimental Procedure 1. Each group will get one sterile 1.5 ml microcentrifuge tube. To it add: a. 200 µl of an overnight culture of E. coli..

b. 100 µl of phage to the cells.

2. Mix this by tapping the tube with your finger to evenly disperse the bacteria and the phage.

3. Incubate at 37oC (not shaking) for 5-10 minutes. During this time the phage infect the

bacteria. 4. During the incubation watch the video entitled “Plating phage”. 5. Obtain a tube of Top Agar from the 55oC temperature block:

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6. Add the bacteria/phage mix to a tube of top agar. Each tube contains 3-4 ml of top agar.

7. Gently swirl the contents of the tube and pour onto an agar plate (rich media),

dispersing evenly. 8. Let the top agar solidify for 15 minutes with the lid slightly ajar to remove

condensation. 9. After 15 minutes close the lid and label the bottom of your agar plate with the

following information: date, section number and group. 10. Your TA will put the plates for the entire class UPSIDE DOWN in a 37oC incubator.

You will examine your plate for plaques and bacterial colonies during Part 2 of this experiment, which will be conducted during the next lab period (Lab 4).

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Name: _________________________

Date: _________________________

TA: __________________________

Section: _______________________ DUE: AT THE END OF LAB LAB ASSIGNMENT #3 - Plasmid, Bacteria and Bacteriophage Week I

1. pUC19 is a commonly used cloning vector. One of the features that makes it useful

for cloning is the presence of several unique restriction sites within the lacZ gene that codes for β-galactosidase.

a. Explain why having more than one unique restriction site is a useful feature

for a cloning vector.

b. Explain why the location of these unique restriction sites (in the lacZ gene) is useful for cloning purposes.

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2. Bacterial strains containing different plasmids are generally indistinguishable under

the microscope. In order to determine which strain contains vector, vector plus insert or lacks a vector, one has to observe growth on media containing antibiotics and indicators, i.e. one must select and screen. Describe in the following chart what media your group chose and what your expectations are for the growth of the different strains on each medium. Recall that no growth on one plate must be balanced by growth on a second plate as a control (this control is to insure that the culture you were given actually had live cells and will grow under appropriate conditions).

Plate Media

E. coli without pUC19

E. coli with pUC19 E. coli with pUC19 plus

insert

3. Growth of phage in the laboratory differs noticeably from growth of bacteria.

a. Why are the phage incubated with bacteria prior to plating?

b. The incubation with bacteria is terminated after 10 minutes. Based on what you know about bacteriophage life cycles, if the incubation was extended to 1 hour would the number of plaques accurately reflect the number of phage in the original culture? Please explain your answer.

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LAB 4 Introduction: During this lab you will complete the experiments begun during Lab 3. Before proceeding, please review the introduction to Labs 3 and 4 (p.24) and the experiments you began last lab session. Grading Lab 4: At the end of lab, each individual will turn in the assignment reporting the results of the restriction digests to distinguish the plasmids with inserts from those without, the identity of the three bacterial strains, and answers to questions regarding the phage cultures.

EXPERIMENT 1 (Part 2) - Restriction Enzyme Analysis of Plasmids Each group will determine which plasmid sample contained the insert and estimate the size of the insert by separating the restriction fragments using electrophoresis on an agarose gel. Experimental Procedure 1. Collect the two restriction digests prepared during Lab 3 from your TA. 2. Spin the digestion tubes in a microcentrifuge (for 30 seconds) to bring down the

condensation. (Be sure to balance the centrifuge before spinning.) 3. Add 2.2 µl of 10x loading dye to each of the two tubes (50% sucrose, 0.2%

bromophenol blue, 0.2% xylene cyanol). The sucrose in the loading dye makes the sample heavy to aid loading the gel, and the blue dyes allow you to visualize how far the samples have run on the gel.

4. Watch the video entitled “Restriction Enzyme Lab” and then practice loading your

sample on the practice gel as shown in the video.

Hints: Slowly and gently aspirate the sample to avoid creating bubbles. Expel any excess air from the pipetter, leaving the sample just at the very tip of the pipetter end ready to be expelled. Insert the pipetter tip into the top of the well and slowly dispense. Once all the liquid is dispensed do not continue to dispense or air bubbles will be introduced into the well.

5. Load the gel after confirming well assignments with your TA (three immediately

adjacent wells per group). Load 15 µl of each digest into separate wells in the gel (one digest per well). Load 15 µl of the standard (or ladder) into the third well. Make sure you note the relative position of the two digests.

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6. Three groups will load on one ten-well gel. Begin electrophoresis after all groups have completed loading the gel.

CAUTION: High electrical voltage is dangerous. Always close the lid of the gel

before turning the voltage on. 7. Carefully slide the lid onto the buffer chamber and run the gel at 100V. Stop the gel

once the dark blue dye is near the bottom of the gel (~60 minutes). 8. Transfer the gel to a Gel Transport Dish. Wearing a glove, CAREFULLY place the

gel into the ethidium bromide (EtBr) bath. Replace the EtBr bath cover and soak the gel for 15-20 minutes.

CAUTION: Ethidium bromide is very toxic – do not touch the gel with your hands and take care not to splash. Wear gloves when handling the gel, remove and dispose

of the gloves immediately after handling the gel. 9. Carefully transfer the gel to the water (dH2O) bath and destain the gel for 5-10

minutes. 10. Transfer the gel to a Gel Transport Dish and take it to the gel documentation system.

Placing it in a dish will avoid dripping buffer onto the floor, which may be contaminated with ethidium bromide.

11. Remove the gel from the dish and place it in the center of the hood on the

transilluminator. Close the door. Turn on the power strip to engage most of the documentation system. After turning on the UV source in the transilluminator (bottom switch on the front), you should be able to view your gel on the monitor. If you encounter problems, consult your TA.

12. Using the print button on the integration control unit, print a copy of the gel. This

print can be cut in thirds to share with the other groups. Make sure each student in your group has a copy of the appropriate third.

13. After you have printed an image of your gel successfully, turn off the UV light in the

transilluminator and the power strip for the documentation system. Discard the agarose gel as directed by your TA. Clean up the transilluminator with paper towel and rinse out the Gel Transport Dish.

14. Gel Intrepretation: Electrophoresis of the molecular weight markers should resolve

10 bands; 10,000 bp, 8000 bp, 6000 bp, 5000 bp, 4000 bp, 3000 bp, 2000 bp, 1500 bp, 1000 bp and 500 bp.

15. For a reminder of how to construct a standard curve using EXCEL see instructions

on p. 12 of lab manual.

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EXPERIMENT 2 (Part 2) - Identification of Bacteria Strains Using

Selection and Screening 1. Collect the plates you streaked during Lab 3 from your TA. 2. Score your plates for growth and colony color. 3. By combining this analysis with the results of your restriction digests, you should be

able to state on the Assignment Sheet which digested plasmid sample (from Experiment 1) is in which bacterial strain (A, B or C).

EXPERIMENT 3 (Part 2) - Growth of Bacteriophage 1. Collect the plates prepared during Lab 3 from your TA. 2. Examine the plates, you should see anywhere from 10 to 100 plaques. 3. Answer the questions on the Assignment Sheet.

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Name: ________________________

Date: _________________________

TA: __________________________

Section: _______________________ DUE: AT THE END OF LAB LAB ASSIGNMENT #4 - Plasmid, Bacteria and Bacteriophage Week 2

A. Plasmid Identification Paste the image of your agarose gel here. Use EXCEL to construct a standard curve and determine the size of each band observed. Label the size of the bands on Figure 1. Please attach your standard curve. Figure 1. Agarose gel documentation of restriction enzyme digest.

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Questions: 1) Which plasmid contains the insert?

2) What size is the insert?

3) What would you have observed, with respect to the size and number of bands, if you had digested the DNA with EcoRI alone? Why?

B. Identification of bacterial strains:

Bacteria Strain

Growth/Color on

LB Plates

Growth/Color on LB + Ampicillin + XG + IPTG Plates

Indicate plasmid contained in strain (none or unknown

code from Experiment 1)

A

B

C

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C. Phage Plaques Examine your plates containing plaques and answer the following questions:

1. a. How many phage from the original phage stock are required to make a single plaque? Why?

b. What was the concentration of bacteriophage (bacteriophage/ml) in the

sample that you plated out?

2. If you had a tube containing a mixture of phage (for example a phage library), what is the advantage of plating out the phage-infected bacteria, as we did in lab, to produce isolated plaques over growing the phage in liquid culture?

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Appendix: Molecular Visualization with RASMOL

RASMOL is a computer graphics program for the visualization of the atomic-resolution, three-dimensional structures of proteins, nucleic acids and small molecules. Using interactive commands at the “RASMOL>” prompt will allow you to look at specific features of the structures. We will use RASMOL to look at the structure of β-galactosidase. The protein structures that are known are deposited in the Protein Data Bank, which is accessible through the world-wide web at www.pdb.rcsb.org/. Getting started: 1. Load the molecule into the program by clicking on the RASWIN icon. 2. On the menu bar, highlight “Open” and from the list of molecules select BGtet.pdb

Displayed is a wireframe model of the β-galactosidase tetramer (recall that the active enzyme is a is four identical polypeptides, each 1023 amino acids in length). A wireframe model shows the positions of all of the atoms (except hydrogen atoms, they are too small to be detected reliably by X-ray crystallography, however we can infer their positions) and covalent bonds. Notice that the oxygen atoms are colored red, the nitrogen atoms blue and the sulfur atoms yellow. You can rotate the molecule by holding down the left mouse button and rolling the mouse. You can zoom in and out by simultaneously holding down the shift key, the left mouse button and moving the mouse toward or away from you. Click on the panel that says “RASMOL COMMAND LINE” found at the bottom of your computer screen. In the window that appears, at the RASMOL> prompt, type show information (press “enter” after all of the commands shown in bold). This gives a summary of the features of the protein structure such as the number of helices, hydrogen bonds, beta strands and turns (there are 8 chains because the crystallographer was unable to view a short region of each monomer, so each monomer is broken into two pieces).

3. Color each monomer differently by typing color chain.

Looking at Secondary and Tertiary Structure: 4. In order to have a better view of the secondary and tertiary structure, type “ribbon”

and press enter. The wireframe atoms of the backbone are hidden by a “ribbon” diagram that emphasizes the shape of the secondary structure elements and how the secondary structure folds together to form the tertiary structure of the protein. The amino acid side chains are still present in wireframe depiction.

5. Type “wireframe off”. Now only the tertiary structure is displayed. Find the α

helices, the β strands.

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Looking at the Five Domains and the Active Site: The 1023 amino acids of each β-galactosidase monomer fold into five compact domains plus ~50 amino acids at the N-terminus which are relatively extended. Focus on one of the four β-galactosidase monomers for the following, or load the monomer pdb file called Bgal.pdb . 6. Domain 1 is a “jelly-roll” type β-barrel. It is formed by 12 β-strands and short

segments of α-helix Type select 50-218, then color red. (50-218 refer to amino acids).

7. Domain 2 is similar in fold to the immunoglobulin constant domains. Select 219-335

then color cyan. The connection (residues 272-288) between the third and fourth strands of this barrel forms a protruding loop that forms part of the active site with the neighboring monomer. Color these residues yellow.

8. To see Domain 3, type select 336-627, then color purple. Domain 3 is a variation on

a fold called the TIM barrel. It contains the catalytic site. You can visualize residues in the active site by typing select 461, 502, 503, 537 then wireframe on, then color yellow.

9. The fourth domain, residues 628-738, is very similar to domain 2. Color it blue. 10. The fifth domain is an 18-stranded anti-parallel β-sandwich. Visualize it by select

740-1020, and then color green. Looking at Hydrogen Bonding: 11. Next, recall that the secondary structure is stabilized by hydrogen bonds between

backbone atoms of the amino acids. To view these for the entire structure, type select all. Then type wireframe on to restore all of the sidechains. Type color cpk to restore coloration by atoms. Then type ribbon off and backbone on, this traces the backbone atoms. Display the hydrogen bonds by typing hbonds on.

Looking at Charged and Hydrophobic Residues: 12. Next, remove the hydrogen bonds by typing hbonds. To display acidic amino acid

sidechains type select all, then select acidic. Color the acidic residues red by typing color red. Display and color the basic amino acids blue by typing select basic then color blue. Notice where in the structure the charged amino acids are found.

13. Now display and color the hydrophobic amino acids by typing select hydrophobic, then color white.

14. Display the van der Waals radii of the atoms by typing select all, then spacefill on

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15. Exit RASMOL by typing exit.