chemistry 3251 biology 3252 biochemistry i 2006 laboratory...

Chemistry 3251 Biology 3252

Biochemistry I

2006

Laboratory Manual

Biochemistry I Lab Manual, Fall 2006 of 35

Table of Contents

Laboratory Safety............................................................................................................ 3 Suggestions for More Efficient Use of Laboratory Facilities ............................................ 6 Laboratory Work and Reports ......................................................................................... 8 Contact information for Biochemistry I........................................................................... 10 LAB #1—Properties and Reactions of Oils.................................................................... 11 LAB #2—Characteristics of Polysaccharides ................................................................ 15 LAB #3—Quantification of gene copy number using southern blotting………………… 19 LAB #4—Protein Analysis: Protein Assays & Protein Denaturation .............................. 30 LABORATORY CHECK IN WILL OCCUR DURING THE WEEK OF SEPTEMBER 11, 2006. YOU MUST CHECK INTO YOUR ASSIGNED LABORATORY THAT WEEK.


Laboratory Safety

A set of departmentally approved general safety regulations is issued, for reading and signature, to all students in chemistry laboratory courses at the beginning of each course. These regulations must be read, understood and adhered to. Safety procedures appropriate to this particular course, some of which may be additional to those referred to above, are listed below. NOTE: Eye protection and lab coats are mandatory for all labs. However, safety glasses, lab coats, and gloves are not provided by the chemistry department. Students must purchase their own safety glasses either from Chem Stores (CB2039) or off-campus, if they prefer. As well, students must purchase their own lab coats, which are available at the bookstore or from off-campus sources. In addition, Chem Stores has reusable nitrile gloves in various sizes for sale. 1) Students are reminded that "Failure to comply with directions of members of the

University administration or of the teaching staff (including laboratory instructors) acting in the proper performance of their particular duties" represents a violation of the Code of Student Behaviour.

2) Any student who disregards safety regulations or becomes a danger to him/herself

or others will be asked to leave the laboratory. 3) Each new experiment is preceded by a brief pre-lab discussion. Please ensure that

you arrive on time because any changes or additional information you require to perform the lab will be discussed at this time.

4) If a lab session is missed for any reason, notify Ainsley or your instructor, however,

no makeup labs will be allowed. 5) No switching of laboratory sections is allowed without written permission from the

instructor in charge of the course. 6) Working in the laboratory in the absence of an instructor is not allowed. 7) No unauthorized experiments are allowed. 8) Make sure you are familiar with the positions and operation of the fire extinguishers,

fire blankets, emergency shower and eye bath, and with the location of the MSDS sheets.

9) Access MSDS sheets prior to the lab period. Make a list of the chemicals used in

the lab in question and outline any hazards, and methods of disposal. 10) Come to lab well prepared. Familiarize yourself with laboratory equipment. Always

read and follow directions carefully. DO NOT TRY TO HIDE MISTAKES!


11) ALL ACCIDENTS MUST BE REPORTED IMMEDIATELY! Any incident which has caused, or which may subsequently cause injury MUST BE REPORTED IMMEDIATELY to the laboratory instructor.

12) Sandals or shorts are not allowed. Lab coats are mandatory and available in the

bookstore. 13) No smoking, eating, or drinking in the lab. 14) Eye protection, by the use of safety glasses is MANDATORY for all persons working

in a chemical laboratory. 15) Gloves should be worn when handling corrosive or carcinogenic materials. 16) Do not work with flammable solvents near an open flame and keep any flammable

solvent in a stoppered flask. Do not heat flammable solvents (e.g. alcohol) with an open flame.

17) NEVER HEAT A CLOSED SYSTEM OF ANY KIND. 18) BOILING CHIPS MUST BE USED WHEN HEATING ALL ORGANIC LIQUIDS. 19) Never use a thermometer as a stirring rod. Report the breakage of any thermometer

immediately. 20) While shaking or venting a separatory funnel, never point a separatory funnel at

yourself or your neighbour. 21) When in doubt about the properties of any chemical, check MSDS sheets or ask the

instructor. 22) Use the fume hood for reactions involving toxic gases and in any case where the

potential hazards of an experiment are unknown. 23) Never taste a chemical and be very cautious about smelling chemicals. If you are

directed to smell a chemical, do so by gently wafting the vapours towards your nose from the top of the container.

24) To insert glass tubing (including thermometers) through a rubber stopper, first

lubricate the tube and stopper with water. Hold the tubing with a cloth near the end to be inserted, and insert gently with a twisting motion.

25) Use a match or a striker to light the Bunsen burner - never paper. 26) Do not use an air jet to dry glassware. Rinse glassware with acetone and dry with

vacuum suction.


27) Do not pipette directly from a reagent bottle. Never pipette liquids by mouth. 28) When diluting a concentrated acid or base, ALWAYS ADD ACID OR BASE TO

WATER. 29) If corrosive liquids touch the skin, flush with water immediately for at least 15

minutes. Have your neighbour/partner notify the instructor immediately. 30) Neutralize acid or base as follows:

Acid on clothing: use quite dilute ammonium hydroxide, or sodium bicarbonate solution. Base on clothing: use dilute acetic acid, followed by ammonium hydroxide or sodium bicarbonate. Acid or base on bench: Use solid sodium carbonate for either followed by water.

31) Broken glass and chemical spills are to be REPORTED and cleaned up immediately

in a safe, appropriate manner. Place broken glassware into the proper waste container. Dispose of all waste in the proper location.

32) As a general rule, water-soluble materials should be flushed down the sink with lots

of water, and water-insoluble solids put in the trash cans. However, for some materials specific disposal instructions will be given by the instructor.

33) NEVER POUR HOT SOLUTIONS OR CONCENTRATED ACID OR BASE INTO

THE ORGANIC WASTE CONTAINERS! 34) Should your experiment catch on fire, turn off the Bunsen burner immediately and

step away. Do not attempt to save your experiment. Call for the instructor immediately.

35) In the event of an emergency, phone Security at Ext. 8911.


Suggestions for More Efficient Use of Laboratory Facilities

The following points will enable you and your colleagues to achieve this aim. Remember, the teaching and technical staff are here to assist you and your co-operation and conduct will be reciprocated. 1. a) You are required to pay a $10 locker key deposit, which will be refunded upon the

return of your key at the end of term. b) Your locker kit should be clean and complete when you receive it. If it is not, see the technical staff immediately. At the end of term, please leave your kit clean for the next student. c) Locker kits are valued at several hundred dollars. Once you have signed out your locker, it and its contents are the responsibility of both you and your partner. It is in your interest, therefore, to lock your locker after use and to ensure you do not leave any items out because they tend to disappear. If, however, you have to leave apparatus or preparations out of your locker, label them clearly; this will ensure they will be there when you return.

2. Do not leave litter or chemical spillages in sinks or on benches or floors. Efficiency

and safety are jeopardized in a dirty laboratory. 3. Do not take reagents from one laboratory to another (unless you know it is the only

supply for that course, e.g. solvents and many organic analytical reagents and test samples, etc.). Always return reagents to their correct location. If a bottle is empty or a particular reagent cannot be found, approach the technical staff.

4. Apparatus put out or set up by the technical staff should not be moved. If for any

reason you think it essential that it be moved, approach the technical staff.

5. Handle all balances with care. Remember, even the more rugged top-loading balances are precision instruments. Points to remember are: a) Do not knock or move the balance; it has been adjusted and leveled for sensitivity and accuracy. b) If you spill materials on the balance, clean them off immediately. Most vapours evolved from solids or liquids will corrode the balance pan and mechanism.

6. You are required to purchase a lab breakage card in order to cover the cost of

broken glassware. You can get this card from Chem stores for a $20 deposit. The card must be obtained before beginning the lab and is valid for all chemistry courses with labs. When a piece of equipment is broken, a set amount is deducted from the value of the card (see the posted price list for amounts). Upon returning the lab card to Chem stores on the completion of all chemistry courses, the $20 deposit will be returned minus any deductions due to breakage. If the full value of the card is consumed through breakage, you must obtain a new card. Please note that lost cards will not be refunded. In the event a card is lost and you still have chem labs to complete, a new card must be obtained.


Before leaving the laboratory, after your experiment is complete, ensure that your work area has been cleaned up and that all of the equipment used has been properly put away, as marks will be deducted accordingly.


Laboratory Work and Reports 1. Four experiments will be performed during the course of the term. Each student will

work with at least one partner. 2. Each experiment will be introduced via a brief pre-lab discussion. Attendance at this

pre-lab discussion is mandatory, since it introduces and explains the necessary techniques and hazards for a given experiment. No student will be allowed to start an experiment if the pre-lab discussion has been missed. No make up labs are allowed. Thus, a mark of zero will be assigned for that lab.

3. Attendance at all laboratory sessions is mandatory. Non-attendance will result in a

mark of 0 for missed labs. Medical or compassionate reasons apply if accompanied by the appropriate documentation.

4. All laboratory sessions are scheduled. Thus, laboratory reports must be submitted

during the laboratory period of the given due date. A penalty of 1 mark per late day will apply. A laboratory report not handed in during the lab period must be handed in before 5:30 on the due date or it will be considered late. After 5 days, a mark of 0 will be assigned.

5. Students are reminded that plagiarism will result in a mark of 0 for all parties

concerned. Although students are encouraged to discuss and share information regarding mechanistic and other theoretical material, each student must submit a completely independent laboratory report in their own words. Identical or similarly worded laboratory reports are unacceptable. Quoting references is unacceptable. Similarly, copying single sentences, entire paragraphs or pages from textbooks is also unacceptable. REFERENCES MUST BE CITED IN THE DOCUMENT AS THEY ARE USED!

6. All laboratory reports must be submitted on stapled, loose-leaf paper. Reports must

be typed or written in ink, on one side of the page only. Each report will include a cover page with title, date, lab section, and name of partner.

7. If a laboratory report cannot be read (illegible or small writing) a mark of zero will be

assigned. Again, typing reports is encouraged. Laboratory report format

Laboratory reports will be written up in the form of a scientific paper. Reports will follow the following format:

a) Abstract—A brief summary of the results you obtained

b) Theory—covers relevant theory of the experiment without discussing specifics of the experiment itself


c) Method—Include any variations from the standard method reported in this manual

d) Results—includes observations, tabular data, graphs, etc. obtained from the experiment

e) Discussion—interpretation and analysis of results; explains WHY things happened and what they mean

f) References—any references used for the lab report. Use references appropriate for the lab material.

A Note Regarding Internet References: The temptation to use the Internet (e.g., Wikipedia) as a reference is understandably great. However, students must learn to identify what is an acceptable reference and what is not. Most websites that contain scientific information are NOT refereed like a journal or textbook. This applies to Wikipedia as well, since the editors are not recognized leaders in their fields.

Acceptable Internet references include:

i) Articles downloaded from a journal publisher’s website. The American Chemical Society as well as several other groups publish their journals on-line as a free resource. Students are strongly encouraged to make use of these excellent resources.

ii) Information taken from a research group’s website. These are identifiable as being hosted by a university or other such institution. Claims made regarding research on a corporate website should not be used.

iii) Technical data and specifications from a manufacturer’s website regarding their products. Looking up the molecular weight of a macromolecule from a chemical supply company is acceptable.

iv) If there are any other references students wish to use, please confirm the source with the professor or lab instructor. Ideally, a link to such a website should be sent via e-mail.


Contact information for Biochemistry I Course instructor: Dr. D. Law [email protected] 343-8277 Office: CB4018 The course website includes the outline, marking scheme and lectures for download. http://flash.lakeheadu.ca/~dlaw/3251.html Dr. Law teaches the course and chose the labs. Lab coordinator and technician: Jarrett Sylvestre [email protected] 343-8540 Office: CB-2049A Jarrett is the main course contact for lab-related information. He sets up, oversees, marks and compiles results for the labs. Lab demonstrators: Chris Edmunds 343-8523 [email protected] Office: CB3006B

David Brookes 343-8735 [email protected] Office: CB3008

Chris and David are involved in lab setup, supervision and marking.


Lab #1 Properties and Reactions of Oils

Introduction Fats and oils (liquid fat) are esters of glycerol and fatty acids. Whether a fat is solid or liquid at a given temperature depends on the nature of the fatty acid. Solid animal fats contain mostly saturated fatty acids, while oils typically contain a high amount of unsaturated fatty acids. Besides the degree of unsaturation, the length of the fatty acid chain also influences whether a fat is solid or liquid. Short chain fatty acids, such as those found in coconut oil, convey liquid consistency in spite of the low unsaturated fatty acid content. The fatty acids most frequently found in vegetable oils are palmitic, stearic, oleic, linoleic, and linolenic acid. Palmitic and stearic acid are saturated fatty acids, while oleic, linoleic, and linolenic acid posses some degree of unsaturation. In this laboratory we will examine some different characteristics of oils in a semi-quantitative fashion. We will also look briefly at one of the most important (at least industrially) reactions involving fatty acides. Part 1: Determining Iodine Number of Oils Fats and oils (liquid fat) are esters of glycerol and fatty acids. Whether a fat is solid or liquid at a given temperature depends on the nature of the fatty acid. Solid animal fats contain mostly saturated fatty acids, while oils typically contain a high amount of unsaturated fatty acids. Besides the degree of unsaturation, the length of the fatty acid chain also influences whether a fat is solid or liquid. Short chain fatty acids, such as those found in coconut oil, convey liquid consistency in spite of the low unsaturated fatty acid content. For various reasons, the degree of unsaturation in oils is an important piece of information. There are a variety of ways to do this, but one of the most common methods is to determine the Iodine Number of a solution containing a triglyceride. Essentially, the iodine number of oil indicates the amount of iodine needed to saturate all the carbon-carbon double bonds in a known amount of triglyceride. By reacting the triglyceride with a known mass of iodine, then back-titrating the excess iodine with sodium thiosulphate, the number of milligrams of iodine needed to saturate a sample can be calculated.


Method Choose one of the oil samples provided and weigh out 1.0 g into a small, clean, dry beaker (50 mL). Dissolve the sample in 4 mL of solvent (the solvent in this case being a 1:1 mixture of ethanol:ether). Transfer the dissolved triglyceride to a 50 mL round bottom flask, making sure to wash the beaker twice with 1 mL of solvent (2 mL total), adding the wash to the round bottom flask each time. Add 25 mL of 0.5 M KOH in ethanol to the distillation flask. WARNING: Potassium hydroxide is extremely corrosive and can cause serious burns on contact with skin!!! Make sure to use appropriate protection during this procedure. Make sure that the potassium hydroxide solution is fresh and free of precipitates. Set up a second round bottom flask with an equal amount of potassium hydroxide solution, plus 7 mL of solvent. This should make the volumes in the two flasks equal (remember, your triglyceride added volume to the initial 4 mL of solvent). Attach a reflux condenser to each flask and heat the solutions in boiling water baths. A large crystallization dish should be adequate for both flasks. Heat the mixtures for 30 minutes. Saponification will occur during this time. Allow the flasks to cool and add 3 drops of indicator solution (phenolphthalein, 10 g/L). The molar difference between the amount of 0.5M HCl required to neutralize the “Control” and the amount of HCl required to neutralize the test sample equals the amount of 0.5M KOH used in the saponification process. Part 2: Determining Saponification Numbers of Oils As triglycerides are comprised of three fatty acids linked to glycerol through an ester linkage, it is possible to hydrolyze that link and produce glycerol and three potassium salts of fatty acids. By heating a triglyceride in aqueous potassium hydroxide, the fatty acid esters can be broken down. The amount of potassium hydroxide required to totally hydrolyze 1 gram of triglyceride is known as the saponification number. Saponification number is related to the hydrocarbon chain length of the fatty acids attached to the glycerol backbone. The longer the hydrocarbon chain portion, the lower the saponification number. Method This method traditionally uses a reagent known as Hanus’ iodine reagent. Although it will be prepared ahead of time by the technician, it is useful to know how much iodine


it contains. Hanus’ solution is produced by dissolving 6.6 g of iodine in 500 mL of glacial acetic acid. To this, 1.6 g of bromine is added. Start by drying 2 500 mL flasks by rinsing them first with ethanol, then with chloroform. Label one flask as being for “Sample” and the other for “Control”. To the “Sample” flask, add a 0.5 g sample of triglyceride. Be sure to measure the precise mass of triglyceride. To each flask add 10 mL of chloroform. Add 25 mL of Hanus’ reagent to each flask, stopper and shake. Place each flask in a dark place (lab locker) for one hour. During this time, the iodine will add to the carbon-carbon double bonds. At the end of the hour, add 50 mL of distilled water to the flasks, using it to rinse the walls and stopper. Add 10 mL of 10% KI solution. This will react with Hanus’ reagent to produce “free” iodine and protons. This free iodine can then be titrated with 0.1 M sodium thiosulphate. At the end point of the titration, the solution will turn a clear straw-yellow colour. At this point, add 1 mL of 1% starch indicator solution. The solution will then turn blue. Continue the titration until the blue colour disappears. Carry out this procedure with both the “Sample” and “Control” flasks. The difference will allow the calculation of how much iodine was bound by the triglyceride. This value, divided by the mass of triglyceride, will yield the Iodine Number. Part 3: Hydrogenation of Fatty Acids One of the most important reactions in industry is the hydrogenation of carbon-carbon double bonds. There are numerous reasons for this, and many different industries that utilize this reaction. There are a number of ways to carry out the reaction, and considerable research has been poured into finding the fastest, most cost effective ways to do so. Many of these involve expensive catalysts, elevated temperatures and pressures. However, in this part of the experiment, a simpler, slower method will be examined. Due to the materials involved, the actual hydrogenation will be a matter of demonstration only. However, analysis of the starting materials and the products will be performed by the students. Method The technician will have assembled a hydrogenation apparatus, consisting of a heating mantle, a three-necked round bottom flask, and a source of hydrogen. The unsaturated fatty acid starting material is dissolved in dichloromethane and mixed with a Raney nickel catalyst. Students will be given a sample of the original starting material.


Using a steam bath, evaporate off the solvent and collect the fatty acid. Use a Fourier transform infrared spectrometer to acquire a spectrum of the fatty acid. The reaction takes several hours to complete, and will have been started well before the start of the lab. The technician will provide a sample of the final product. Again, place the sample in a small beaker with boiling chips and evaporate off the solvent with a steam bath. Collect the crystals from the bottom of the beaker and prepare an IR spectrum. Note the differences between the spectra of the starting material and the product. Explain these differences. Waste Disposal Pour liquid waste into appropriate containers in the fumehoods. The technician and demonstrators can also point out which solutions can safely be poured down the drain.


Lab #2 Characteristics of Polysaccharides

Introduction Carbohydrates are polyhydroxyl compounds, which usually have an aldehyde or ketone functional group, although it is commonly in a hemiacetal or acetal form. The simplest carbohydrates are known as sugars or saccharides. These units can then link together and form extensive biopolymers. Depending on the arrangement of these saccharide units, the polymer will have different properties. Common polysaccharides in nature include cellulose and a wide variety of starches. They form the building blocks of plant matter and have found widespread usage in almost every application of day to day life. In this experiment, we will explore some of the reactions polysaccharides can undergo. Some of these reactions show how the polymers can be altered for specific applications. Others show parallels of how these biomolecules are used in their natural state. Part 1: Synthesis of Cellulose Acetate from Cotton The chemistry of polysaccharides is essentially the same as the chemistry of alcohols. The many hydroxyl groups on these polymers can undergo many of the same reactions that simple alcohols are subjected to. One of these reactions is esterification, a chemical addition of an acid to an alcohol. Cellulose is a very common biopolymer found in certain plants. Industrially, humans have been using cellulose for millennia for a variety of uses. However, it has only been in the last hundred and fifty years or so that it has been chemically altered to serve even more purposes. Acetylation involves the addition of an acetyl group (provided by acetic acid and acetic anhydride) to the hydroxyl groups on the individual monomer units of the polysaccharide. The resulting polymer is still structurally similar to cellulose, but has some very different properties. Method Combine, in a 50 mL Erlenmeyer flask, 7.5 mL of glacial acetic acid and 5.0 mL of acetic anhydride. Soak 0.15 g of cotton in distilled water for one minute. Squeeze it dry with fingers, then press between paper towels to blot out as much moisture as possible. Gently pull the fibres apart (note: Do not ‘shred’ the cotton, as your product


will be too fine to precipitate out) and place in the flask. Use a glass stir rod to press the cotton down below the surface of the solution. Add 3 drops of concentrated sulphuric acid. The solution should warm, but not get hot. If it becomes hot, there was too much water in the cotton, and you won’t get the desired product (why?). Swirl the flask and leave it to stand for 10 to 15 minutes. At the end of that time, the cotton should be completely dissolved. Clamp the flask in a hot water bath or over a steam bath, prepared earlier. Allow it to heat for 10 to 15 minutes with occasional stirring. This should dissolve the last of the cotton. If any cotton remains, add a few drops of acetic anhydride and continue heating until the solution is homogeneous. Remove from the heat and allow the flask to cool to room temperature. Then place the flask in ice for one minute. Pour the contents of the flask into an ice water mixture (20 mL of water and 20 g of ice in a 250 mL beaker) slowly, while stirring gently (vigorous stirring will cause the product to be too fine to filter out). After 10 minutes, the cellulose acetate should precipitate out as a gelatinous mass. Vacuum filter the mixture using a Buchner funnel, and collect the precipitate. Rinse the product twice with 10 mL portions of cold distilled water. Leaving the cake of product on the vacuum, pat it down occasionally to speed drying. Allow to dry for 10 to 30 minutes. If after 10 to 15 minutes on the vacuum, the product is still a wet paste, you’ll need to press it dry between paper towels. Once the product is dry enough to crumble, place it in a clean, dry 50 mL Erlenmeyer flask with 10 mL of dichloromethane. If it doesn’t all dissolve, add a few more mLs of dichloromethane. If the solution is cloudy, dry it with a small amount of anhydrous sodium sulphate. Decant the solution into a 50 mL beaker and add 3 drops of diethyl adipate. Place the beaker in a water bath at a temperature of 40 C and evaporate off the solvent. Be sure to remove the beaker before all the solvent is removed! You will achieve a better quality of film if you heat it slowly and gently. If heated too fast or too vigorously the film will crack and decompose. Allow the rest of the dichloromethane to evaporate and show the film to the lab instructor. Then test the solubility of the cellulose acetate in water by adding a few drops of distilled water to the beaker. This will also allow you to lift the film out of the bottom of the beaker. For comparison purposes, test the solubility of cellulose in dichloromethane by adding 3 mL of solvent to a test tube and then adding a small wad cotton. Record all your observations. Part 2: Hydrolysis of Starch by Enzymes and Acid Starch is a polysaccharide consisting of repeating glucose units. These are linked together by α (1 4) linkages. Branching of the starch molecule can occur by the


random formation of α (1 6) bonds. When starch is broken down, it is broken down into progressively shorter and shorter chains. The enzyme alpha-amylase (found in human saliva) breaks down starch into chains of random length. Beta-amylase is excreted by the pancreas, and it cleaves starch residues into maltose (two glucose units), maltotriose (three glucose units) and dextrin (branched residues). Starch can be hydrolyzed by acid or through the use of these amylase enzymes. Both are effective, but they each require different conditions to achieve the same results. The following method will show that the conditions for one are much harsher than the other, and will demonstrate just how effective enzymes are at catalyzing reactions. Method This is a spectroscopic method that uses the absorption of a solution at 570 nm to determine concentration of starch remaining. Before starting this part of the experiment, set up a UV-Visible spectrophotometer, set the wavelength of absorption to 570 nm and allow it to warm up. Zero the reading with a cuvette of distilled water. This procedure will use stock solutions of starch, enzyme, acid, and iodine. The stock starch solutions will need to be diluted. The graphs will require 8 different concentrations. The first (0.01 mg/mL) and the last (0.15 mg/mL) have been provided, along with two others (0.05 and 0.11 mg/mL). Students will be responsible for calculating the dilution factors needed to generate the other four points themselves. Students will need approximately 25 test tubes per group, as they will be performing three different ‘runs’, each consisting of 8 tubes. To each of the eight tubes, add 8 mL of the different concentrations. Prepare the different ‘runs’ as follows: 1) To generate a standard curve, add 1 mL of distilled water to each of the eight tubes, and place them in the incubator for 10 minutes. At the end of the 10 minutes, remove the test tubes and cool them in ice. Add 1 mL of iodine solution, and measure the absorbance of the solutions at 570 nm. This will give a ‘baseline’ of the response of known concentrations of starch to the iodine solution. 2) To see how quickly the enzyme-catalyzed reaction proceeds, add 1 mL of the stock (0.4 mg/mL) alpha-amylase solution to each of the eight starch samples. Place them in the incubator at 37 C for 10 minutes. At the end of the 10 minutes, remove the test tubes and cool them in ice to halt the reaction. Add 1 mL of iodine solution to each tube, and measure the absorbance of each solution at 570 nm. Not only will this give a new curve, but the change in concentration of starch can be calculated for each point.


3) To see what conditions are necessary for the acid hydrolysis to proceed, add 1 mL of 0.2 M hydrochloric acid to each of the eight starch samples. Place these in a beaker of boiling water for three minutes. At the end of the three minutes, remove the test tubes and allow them to cool to room temperature. The reaction will actually stop as soon as the tubes cool slightly. Measure the absorbance of each solution at 570 nm. These values can show how much the concentration has changed under these conditions, and allow a comparison to the enzyme-catalyzed reaction. Calculations Use the standard curve to generate an equation for the line of best fit. This equation can then be used to determine the actual concentration from the absorbance measurements. Use this to determine the concentration (in mg/mL) of starch at each point on the other graphs. The rate of consumption can be found by calculating the change in concentration at any point on the graph, divided by the time of reaction (10 minutes for the enzyme-catalyzed reaction or 3 minutes for the acid hydrolysis). Make sure to compare the rates in the Discussion portion of the lab report and consider the relative concentrations of the reagents. Waste Disposal Dispose of all test solutions in the appropriate waste containers in the fume hoods. Ensure that rinsing of equipment containing test solutions are also poured into the appropriate waste containers. Dispose of excess sugar solutions by pouring them down the drain and rinsing with excess water.


Lab #3 Quantification of Gene Copy Number Using Southern Blotting

Introduction The application of molecular biology techniques for the analysis of complex genomes by analysis of the DNA sequences that they contain depends heavily on researchers’ ability to prepare pure, high-molecular-weight, intact genomic DNA. Purification of genomic DNA involves two steps:

• cell lysis and DNA solubilization, followed by • one of several enzymatic or chemical methods to remove contaminating

proteins, RNA, and other macromolecules. The basic approaches described here for plant leaf tissue are generally applicable to a wide variety of starting tissues, including those from bacteria, fungi and animals. Though a molecule of DNA differs in sequence between species, its basic chemical properties are identical in all organisms, allowing for this standardization of procedures. Gel electrophoresis is the fractionation method of choice for nucleic acids. In the following discussion of electrophoresis, it is extremely important to remember that lethally high voltages are often used. Because of the potential hazard, we use an adequately shielded apparatus, an appropriately grounded power supply, and most importantly, common sense, when carrying out electrophoresis experiments. Empirical studies have identified several important variables that affect migration of nucleic acids on gels, including the conformation of the nucleic acid, the pore size of the gel, the voltage gradient applied, and the salt concentration of the buffer. The most basic of these variables is the pore size of the gel, which dictates the size of the fragments that can be resolved. In practice, this means that larger pore agarose gels like those we will use in this lab are used to resolve (or physically separate so that they are visibly distinct) fragments larger than ~100 bp, and smaller pore acrylamide gels are used for smaller fragments. The pore size is varied by changing the concentration of the agarose that makes up the solid matrix of these gels: the higher the agarose concentration, the smaller the pores. This increases the resolving power of the gel for DNA fragments of various sizes (see table below [from Ausubel et al. 1989]). We will use a 0.7% (w/v) agarose gel as a good compromise between the resolution we desire and the fragment sizes we hope to visualize.


Agarose (% w/v)

DNA fragment resolution range (kb)

0.5 30 to 1 0.7 12 to 0.8 1.0 10 to 0.5 1.2 7 to 0.4 1.5 3 to 0.2 Frequently, it is desirable to identify an individual fragment in a complex mixture that has been resolved by agarose gel electrophoresis. This is accomplished by a technique called southern blotting, in which the fragments are transferred from the gel to a filter and the fragment of interest is identified by hybridization with a labeled DNA probe. This probe has traditionally incorporated a radioactive isotope, usually the powerful beta (electron) emitter 32P, to enable it to be visualized on the blot using X-ray film. The energy from 32P is sufficient to cause irreversible damage to cellular macromolecules, and it is thus not suitable for use in teaching labs. However, there are now a number of nonisotopic methods used to make probes. You are not responsible for knowing the details of probe construction, but briefly, we made probes for two genes using a modified nucleoside. Antibodies are used to recognize this modification. The antibodies are covalently ligated to an enzyme that converts a colorless substrate into one that emits light. Thus, when the probe is hybridized to its DNA target on the southern blot, antibodies can recognize it. The location of the probes hybridized to their DNA targets can then be detected by placing the blot into a lightproof box and using an imaging system to detect the light-emitting bands. Southern blotting is widely used to identify specific fragments in a digest of total genomic DNA. In our case, you are using this technique to probe the structure of maize genomic DNA. Specifically, you are interested in determining how many copies of two genes are present in maize DNA:

• Gamma-zein (a storage protein expressed in maize seed) • Ubiquitin (a small protein involved in protein turnover in the cell)

On your southern blot, you will include intact (undigested) genomic DNA, and DNA cleaved into smaller fragments by sequence specific endonucleases called restriction enzymes. By comparing the pattern of bands on southern blots that include DNA from undigested and digested maize DNA, you will be able to (roughly) determine the number of times both of these genes appear in the maize genome (a/k/a gene copy


number). Note that there are more exact ways to determine this information, but that these require expensive instrumentation and technical support. Southern blotting remains a useful technique for (relatively) quick determination of copy number. Why is this important? Multiple copies of genes present in a genome infer that different forms of the gene may be expressed under different environmental conditions. The distinct proteins that result from transcription and translation of these genes may differ only slightly at the amino acid sequence level but have distinct functions in the cell. These are called protein isoforms. The distinct genes coding for the same protein are very likely under the control of promoters that are regulated differently. Thus, one isoform favoring maintenance of metabolism under, for example, heat stress might be present in the cell at high temperatures, while another isoform induced by cold stress is present at low temperatures. This ability to fine-tune metabolism is critical to ensure the organism’s survival. How will you determine the copy number for each gene? We can do some math to figure this out. Both restriction enzymes you will use (EcoRI and HindIII) cleave DNA at 6-nucleotide long recognition sequences. These sequences can be looked up using web tools like those at New England Biolabs: http://www.neb.com/nebecomm/EnzymeFinderSearchByName.asp. First, assume randomness of distribution of the four nucleotides in the genome (a dangerous assumption, but one that we will use). The probability that any particular nucleotide will be present at a given locus in the genome is 1 in 4 (e.g., 1 in 41). A restriction enzyme having a one-nucleotide recognition sequence would thus yield an average fragment length of one nucleotide. Similarly, a particular 4-bp sequence will occur once every 44 bp = 256 bp, while the sequence recognized by “six-cutter” enzymes cuts such as EcoRI and HindIII will occur once every 46 bp = 4.1 kb. These enzymes would generate restriction fragments with an average length around these values. In the model plant Arabidopsis, the average gene size is 2 kb (http://cnx.org/content/m11317/latest/). There is some comparable information available for maize that indicates similar gene sizes exist in this species, but the genome will have to be sequenced to determine this. Thus, on average, restriction digestion with six-cutters should yield one or two bands on southern blots if a single copy of a gene is present in the genome. This means that the probability of the enzyme cutting inside of the gene is relatively low. By contrast, if more than one copy of the gene is present, multiple bands would be expected because restriction


digestion should generate multiple fragments of DNA that possess a copy of the gene (or part of the sequence). To visualize this, picture a length of genomic DNA: Since the size of the gene is small relative to the size of the restriction fragments generated, the frequency of restriction is low. One would expect one or two (because of the 2 kb/4 kb ratio, there is a ~50% chance that there is a single restriction site inside any given gene sequence) bands on a southern blot for EcoRI digested DNA. How many bands would be expected on a southern blot from the restricted DNA shown in the example above? By contrast, if multiple gene copies are present, the probe will hybridize to each one of these. Since each copy is in a different part of the genome (though some may be physically closer than others), and each genomic DNA fragment carrying each of (or a piece of each of) these genes is of a different size, it would be expected that the number of bands on a southern blot increases as the gene copy number increases. In the example above, there are 2 copies of the gene. How many bands would you expect to see on a southern blot of genomic DNA digested with EcoRI as pictured above? Thus, southern blotting is very useful for probing how gene expression is potentially regulated in organisms. New forms of the gene may have arisen through duplication and mutation of existing genes; many of these are present on the same chromosome

Average gene size ~ 2 kb

Average length between restriction sites ~ 4 kb

EcoRI site EcoRI site Gene specific probe hybridizing to gene target

Small section of a chromosome

(made of genomic DNA)

EcoRI site EcoRI site EcoRI site EcoRI site

Gene copy #1 Gene copy #2

Intergenic region of varying length


(as in the example above). If present on a different chromosome, the new copy may have arisen by genetic rearrangement or transposition from a pathogen’s genome . If multiple forms of a gene are present in the organism’s genome, this might suggest that the metabolic pathways containing the protein encoded by these genes is subject to transcriptional regulation. This means that the activity of the pathway’s proteins is largely controlled by the level of their mRNAs. This indicates that differentially expressed genes encoding functionally distinct protein isoforms might be more adapted to certain environmental conditions. By pinpointing the small sequence differences between gene copies, new probes can then be designed to specifically target the environmentally adapted gene and its products. In this way, its function can be characterized during stress and recovery, or during a specific developmental stage. If expression of this form of the gene can be enhanced, the plant might become more resistant to stress. Prior to going to all of this effort, the importance of the gene product (protein) should be demonstrated using traditional biochemical methods. Methods This is a 3 week lab, with specific procedures to be done each week. Even so, there are many steps in the protocol that must be performed outside of the lab timeframe. The demonstrators will perform these tasks for you, and they are indicated in the table below. This is a good indication that performing southern blotting is a time consuming task best accomplished over several days. This also indicates how many molecular biology protocols do not require a large time investment all at once but provide many potential stopping places. Many steps can be performed overnight or over several hours, freeing you for other lab tasks. We will be taking advantage of this multitasking potential by putting you to work on several things at once in this lab.

Performed by

Task list for southern blotting (in order) Week

TA Class 1. Extract maize leaf genomic DNA 1 X 2. Digest DNA with restriction enzymes 1 Finish Start 3. Pour agarose gel 2 X 4. Run agarose gel 2 X 5. Equilibrate gel and set up transfer to blot 2 X 6. Remove blot from sandwich and store 2 X 7. Prehybridize blot 3 X 8. Add probe and hybridize with DNA on blot overnight

3 X


9. Wash blot 3 X 10. Develop and visualize DNA bands 3 X 11. Interpret data and write report post-

3 X

Week 1: DNA isolation and restriction digestion We will use young (10-day-old) maize leaves as a source of genomic DNA, and isolate it using a simple method that works well for leaf tissue. Each step in the method serves to eliminate one form of macromolecule from the crude extract until all that is left is pure genomic DNA. More expensive kit-based methods are available, but note that many kits do not perform as advertised and that a good biochemist often has to tweak kit protocols or abandon them altogether and use a non-kit (a. k. a. “hombrew”) method to isolate the biomacromolecule that they want to study. 1. Remove a pair of flag leaves from a 10-day-old maize seedling. These will weigh

approximately 200 mg each. Place it into a mortar. 2. Add 500 µl of Genomic DNA extraction buffer. Using a pestle, grind the tissue

as finely as possible. This process breaks down the tissue and liberates the DNA. Add another 500 µl and repeat. Transfer the contents to a test tube by pouring. Rinse the mortar with another 400 ul of extraction buffer, and add to the test tube.

3. In a fumehood, add an equal volume (1.4 ml) of chloroform:phenol (1:1). The

phenol must be buffered to a pH=8.0. Mix the chloroform:phenol before adding it. Mix the contents of the tube by flicking tube and vortexing. It should be mixed vigorously to make sure that all the DNA partitions into the aqueous layer.

4. Spin the balanced tube at 40 speed in the clinical centrifuge for 10 min. 5. Remove the upper aqueous layer without disturbing the white precipitate at the

interface using a disposable Pasteur pipette. The protein and cell debris partitions into the lower (chloroform organic) green layer while the DNA dissolves in the aqueous buffer layer. Transfer the upper layer into a clean, dry tube.

6. Add an equal volume of isopropanol (1.4 ml). Mix gently until thread-like strands

of DNA form a visible mass. The best method for this is to add the alcohol with a micropipetter down the inner walls of the tube. The interface between alcohol and aqueous layers will turn cloudy as you gently swirl the contents. The threads of DNA will be visible almost immediately as the green colour of the lower layer


vanishes. Let sit 5 min at room temperature, then centrifuge 5 min as above. The DNA is now (loosely) pelleted at the bottom of the tube. Some of the DNA will adhere to the walls of the tube as a white film, which is acceptable as well.

7. Carefully decant the supernatant by pouring it off onto a paper towel. As you

pour, hold the pellet on the top of the tube and watch it to make sure you do not lose it. Blot open tube top against clean sections of paper towel to absorb as much of the supernatant as possible (tapping carefully to facilitate this).

8. Dissolve the pellet in 1 ml TE buffer by flicking. If there is any debris in the tube

after this, centrifuge 5 min and transfer the supernatant to a new tube. Otherwise, continue to step 9 with the DNA/buffer solution.

9. Add 0.1 volume of 3-M sodium acetate and 2 volume of ethanol (0.1 ml and 2 ml,

respectively). Mix to re-precipitate the DNA as before. Centrifuge, decant and blot the tube.

10. Wash the DNA with 2 ml of room temperature 70% ethanol and centrifuge as

above. 11. Carefully decant the supernatant and dry the tube as above. The DNA pellet is

very loose at this point and care must be used to avoid pouring off the pellet onto the paper towel.

12. Air-dry the pellet in the tube for 5-15 minutes in the fumehood. With the tube

resting in a test tube rack, there should be no ethanol left pooling in the bottom of the tube after this period.

13. Rehydrate the pellet in 200 ul TE buffer. Add the buffer and allow to sit for 15

minutes. Do NOT shake it during this period. At the end of 15 minutes, the pellet should have dissolved completely in the TE buffer. If some DNA remains, swirl the tube gently to complete the process. During the 15 minute waiting period, turn on a UV/Visible spectrophotometer and set it to 260 nm to allow its UV bulb to warm up. Also use this time to set up your restriction digests (see below).

14. Quantify your DNA. Use a quartz or UV-transparent disposable

semimicrocuvette. Blank first with 3 mL water, then add 15 µl of your DNA, invert 3 times to mix and read the A260. Convert the absorbance reading to µg of DNA using the following equation:


A260 x 0.05 x dilution factor = mg/ml DNA in your preparation

For example, if your A260 reading was 0.2 and you added 15 µl DNA to 3 mL water, your dilution factor would be 3000/15 = 200, and the DNA concentration would be 0.2 x 0.05 x 200 = 2 mg/ml. Write the DNA concentration on the tube (and in your notes).

15. For the restriction digests of your genomic DNA, each group will have 3 tubes: one

control (undigested, without restriction enzyme added), and one each of DNA to be digested by EcoRI or HindIII. Each tube should contain 5 µg of DNA. Use the table below to plan your restriction digests (and be sure to present your finished table in the Materials and Methods section of your report). When adding the materials to your tubes, make sure to use the correct technique due to the small volumes used. Place the tip of the micropipette at the bottom of the tube when adding liquids. The best thing to do is to add the distilled water to the tubes first, as this will make it easier to add 1-2 µL amounts if there’s already liquid present. Note: Most store-bought enzyme buffers (including the ones used in this lab) have BSA added. What is the function of the BSA (bovine serum albumin)? Add 2 microlitres of the buffer. To the control tube, add 2 µL of EcoRI buffer.

Component Control

(µl) Eco I digest (µl)

Hind III digest (µl)

5 µg DNA (at _____ mg/ml)

10 x enzyme buffer Distilled water (to 20 µl) Enzyme (add last) 0 1 (EcoRI) 1 (HindIII) TOTAL VOLUME 20 20 20 16. Cap and flick the tubes to mix. Shake down contents of tubes to bottom. Place

them in a small beaker and incubate your 3 tubes at 37°C overnight. The next morning, the demonstrator will remove the tubes from the oven and store them at 4°C until the next lab session.

Week 2: Separation of restriction fragments by agarose gel electrophoresis and blot setup This week you will separate the products of your digestions using agarose gel electrophoresis. The gels have been prepared ahead of time by your demonstrators.


Once the electrophoresis is complete, you will then check the quality of your genomic DNA and restriction digestions using UV light, then set up the transfer of your DNA fragments to a nylon membrane (the southern blot). This is the next step in identification of your target genes within the maize genome.

1. Each lane on the gel will contain the same amount of DNA (how many µg?). To your restriction digestions, add 5 µl of 6X DNA gel loading dye. Make sure the loading dye is added to the liquid at the bottom of the tube. Mix by flicking, and spin tube contents down to bottom by briefly centrifuging if necessary.

2. Load gels using a micropipettor. Your demonstrator will show you the technique. The leftmost lane on each gel will contain molecular weight standards. Load 5 µL of the premixed standard. This mixture of DNA fragments of known size will be used to size the bands you will obtain on the southern blots. Be sure to mark what lanes contain which samples!

3. DNA bands will be electrophoresed towards the cathode (positive electrode). Why is this? Make sure that you have the electrode leads plugged into the appropriate plugs on the power supply. Run the gel at 100 V for ~45 minutes. Early during the run, the blue loading dye will resolve into an upper and lower band. These are two different dyes. Let the lower dye (bromophenol blue) run about 2/3 of the way down the gel, then stop the electrophoresis run by shutting off the power supply.

4. Remove the gel carefully from the gel box and place in a glass tray. Visualize the digestion products and molecular weight standards using UV light, and document the major digestion product sizes relative to the standards. Make note of the positions of the molecular weight standards by using the UV-fluorescent gradations on the gel dock. Calculate their Rf values. Plot a standard curve of Mr versus log(distance migrated in cm). Note the slope and use this equation to calculate the sizes of major digestion products on your gel. This equation will also be used in part 3 to calculate the sizes of the fragments of genomic DNA hybridizing to your probe on the southern blot.

5. The gel must now be treated to denature the double stranded genomic DNA into single stranded DNA that can hybridize to the probe. In its glass tray, treat the gel, sequentially, with:

• 200 ml of 0.2 N HCl: 10 min, decant • Water: 5 x 3 min, decant


6. Measure gel. Cut nylon membrane, 10 pieces of thick filter paper (“3MM”) and a ~7-cm stack of paper towels to this size. Set up an overnight transfer as a gel “sandwich” as shown in the figure. The tray contains transfer buffer (0.4-M NaOH plus 0.6-M NaCl). The nylon filter is first completely wetted in distilled water and then soaked in transfer buffer. The nylon filter is overlaid with one piece of transfer buffer-soaked 3MM, followed by the stack of dry 3MM and dry paper towels. Allow transfer to proceed overnight. The nylon membrane is positively charged; can you explain why?

7. The next morning, the demonstrator will disassemble the sandwich, remove the blot and mark the positions of the lanes with a pencil. The air-dried blot will be rinsed briefly (5 min) in a salt solution (2 x SSC buffer) and stored at room temperature until next week.

Week 3: Prehybridization, hybridization and detection of probes

1. The night before the lab, the demonstrator will equilibrate the buffer for one hour in hybridization solution. This is called prehybridization, and ensures that only DNA on the blot possessing the complimentary sequence to the probe will bind to it. Without prehybridization, the probe would stick all over the blot. Prehybrization occurs at the temperature used to anneal the probe to the blot during hybridization. This is generally 42°C. Pre- and hybridization involve the use of a hybridization oven, where the blots rotate in tubes that continually bathe them in a small volume of the solutions.

2. After a 1-hour prehybridization, the probe was denatured 2 min at 100°C, cooled rapidly on ice, and 0.5 µl probe added per ml of hybridization solution used. (We will use ~5 ml hybrization solution per blot, so we will add 2.5 ml probe to each.) The prehybridization solutions were discarded, and hybridization solutions added. Half of the groups in the class had their DNA hybridized to the zein probe, the other half to the ubiquitin probe. You can then share information for your writeups. The blots were hybridized overnight at 42°C.

3. Wash the blots. Note that the salt concentration decreases and wash temperature increases as the washes progress. This is known as increasing the stringency of the washes. Lower salt and higher temperature are used to progressively remove nonspecifically bound probe from the blot, leaving only probe-target hybrids. Wash the blot as follows to remove unbound probe (~50 ml per wash):

• Blot Wash #1: 2 x 5 min at room temp (RT).


• Blot Wash #2: 2 x 15 min at 55° C. • Buffer 1: briefly (1 minute) at RT.

4. Incubate with antibody-conjugate for 30 minutes (save at 4° C). This antibody binds to the modified base used during probe synthesis, allowing its detection because the antibody is bound to an enzyme.

5. Remove unbound antibody-conjugate by washing 2 x 15 minutes with Buffer 1.

6. Equilibrate blot with ~50 ml Buffer 3. The probe-target DNA bands are now ready to be visualized.

7. In a test tube, add 3 µl CDP-Star to 3 ml Buffer 3. The CDP-Star is a colorless substrate that is converted to a form that emits light by the enzyme conjugated to the antibody (in other words, the substrate is converted to a chemiluminescent product). Mix by vortexing gently.

8. Transfer blot carefully from Buffer 3 by holding it by a corner with foreceps, then letting it drip by a corner for ~5 seconds back into the tray. Gently drag the blot’s lower edge along the tray edge to fully remove excess buffer.

9. When ready to detect the bands (but not before!), transfer the blot face-up to a plastic divider sheet. Pipette the CDP-Star solution carefully directly onto the blot (do not touch the blot with the pipette tip), and overlay with another plastic sheet. Smooth out bubbles.

10. Place the blot in the imager, and take a series of pictures to visualize the chemiluminescent bands. Exposures may need to be varied between seconds and minutes to optimize the signal (probe-target hybrids) to noise (nonspecific probe binding all over the blot) ratio.

Save a copy of the blot on a memory stick so that you can incorporate the picture and information into your report.


Lab #4 Protein Analysis: Protein Assays & Protein Denaturation

Introduction Proteins are biopolymers responsible for nearly all biological functions. Many are enzymes or biological catalysts that are responsible for the chemistry of living organisms. Proteins are mainly composed of α-amino acids, yet there exist many thousands of different molecules. Among many proteins the chemical differences are small or subtle, yet these differences may result in remarkable differences in biochemical function. Proteins in their natural 3-dimensional form or tertiary structure are in their “native state”. Any change in structure from a protein’s native state is called denaturation and often results in decreased or no activity. These changes occur in the secondary and tertiary structures of proteins. In this experiment we will examine a few of the many means of denaturing a protein, specifically phycobiliproteins. Phycobiliproteins are brightly coloured and highly fluorescent proteins found in cyanobacteria (a. k. a. “blue-green algae”, such as Spirulina species, which are often used as nutritional supplements). Spirulina cells contain two easily extractable phycobiliproteins that are dark blue in colour with a strong absorption at 625 nm; they also exhibit a red fluorescence. If denatured, the proteins will loose both their red fluorescence and dark blue colour. This allows for denaturation of the protein to be easily observed. This denaturation can be studied both qualitatively (by examining the colour and fluorescence of the solution) and quantitatively (by examining the absorption at 625 nm). In biochemical study and research, it is often desirable and necessary to determine the protein concentration of a solution. Many quantitative assays have been developed, including the Kjeldahl assay, UV absorption of tyrosine and tryptophan residues, and the Lowry assay. In this experiment we will examine the Bicin-Conicin assay and the Bradford assay; the latter being the most commonly used protein assay. Method Part 1: Qualitative Study of Protein Denaturation For a qualitative study, we will examine protein denaturation via a variety of methods. Isolate the phycobiliproteins using the following method:


Tare a sheet of weighing paper on the balance. Weigh out 750 mg of the powdered Spirulina. Transfer the Spirulina to a mortar and add an equal mass of fine silica. Grind the contents to a fine homogeneous mixture (approximately 5 minutes). Maximize the breaking of the cell walls by grinding the contents against the sides and bottom of the mortar. Divide the contents of the mortar evenly between two 50 mL centrifuge tubes. Add 25 mL of 0.1 M potassium phosphate buffer (pH 7) to each centrifuge tube. Mix thoroughly with a glass rod. The solution should turn a deep blue colour. Balance the centrifuge tubes to within 0.1 g of each other and centrifuge at high speed for 10 minutes. After centrifugation, decant the liquid into a small beaker. Gravity filter this solution into another small beaker and discard the precipitate. The resulting solution of phycobiliproteins should be dark blue in colour and exhibit red fluorescence when held close to a light source (lamp bulb or flashlight). Examine the denaturation of the protein using the following methods. Denaturation can be observed by a loss of fluorescence, change in colour, or precipitation. Note and describe the changes for each method and tabulate your results. Effects of pH In each of two test tubes, place approximately 1 mL of protein solution. To one, add 1-2 mL of 6 M HCl. Repeat with 6 M NaOH Heavy Metal Cations In each of two test tubes, place approximately 1 mL of the protein solution. To one, add 1-2 mL of 0.2 M copper (II) sulphate drop by drop. Repeat with 0.2 M lead (II) acetate. Precipitating Acids In a test tube place approximately 1 mL of protein solution. Add an aqueous solution of 10% (w/v) trichloroacetic acid (Caution: extremely corrosive!) drop by drop. Mix thoroughly. Organic Solvents To 1 mL of the protein solution add 2-3 mL of acetone. Mix thoroughly. Repeat the test with aqueous solutions of 25% (v/v) ethanol, 50% (v/v) ethanol, and 80% (v/v) ethanol. Urea In a test tube place approximately 1 mL of protein solution. Add 3 mL of 8 M urea. Mix thoroughly.


Temperature In a test tube place approximately 1 mL of protein solution. Add 2 mL of 0.1 M potassium phosphate buffer (pH 7). Place the test tube in a boiling water bath for 5 minutes. Part 2: Quantitative Study of Protein Denaturation For a quantitative study, we will examine protein denaturation via the addition of an organic solvent (ethanol) and thermal denaturation (heat). Isolate the phycobiliproteins in the following manner: Tare a sheet of weighing paper on the balance. Weigh out 2 g of powdered Spirulina. Transfer the Spirulina to a mortar and add one half the mass of fine silica. Grind the contents of the mortar to a fine homogeneous mixture (approximately 5 minutes). Maximize the breaking of the cell walls by grinding the contents against the sides and bottom of the mortar. Transfer the powder to a small beaker and add 50 mL of 0.1 M potassium phosphate buffer (pH 7). Mix thoroughly with a glass rod for several minutes. Transfer the mixture evenly into two centrifuge tubes and balance the tubes to within 0.1 g of each other. Centrifuge at high speed for 10 minutes. After centrifugation, decant the liquid and gravity filter into a small beaker. To the filtrate add potassium phosphate buffer solution until the final volume is approximately 100 mL. Precipitate the phycobiliproteins by adding 29.5 g ammonium sulphate. The solution is now a 50% (w/v) saturated ammonium sulphate solution. Add a stir bar and mix at a low speed for 15 minutes. Transfer the mixture evenly into four centrifuge tubes and balance the tubes to within 0.1 g of each other. Centrifuge at high speed for ten minutes to pellet the precipitated protein. Discard the supernatant. Resuspend the pellet in 25 mL potassium phosphate buffer by successive mixing and transfers to each centrifuge tube. In between transferring the solution to each tube be sure to mix thoroughly for a couple of minutes in order to dissolve as much of the protein as possible. After mixing the solution in the final centrifuge tube, gravity filter and collect the filtrate in a small beaker. This filtrate is your final phycobiliprotein solution. Test the absorbance of the protein solution at 625 nm. If the absorbance is not between 0.5 and 0.9, not including absorption due to the cuvette, dilute accordingly. To complete the three denaturation studies, you will need a total of 35-40 mL of protein solution after dilution.


Organic Solvent—Addition of Ethanol To five test tubes, add 2 mL of your protein solution. Before each transfer, swirl the protein solution in order to evenly suspend any undissolved material. To these test tubes will be added the following combinations of ethanol and potassium phosphate buffer:

(i) 0% ethanol, 100% buffer (2 mL total volume added) (ii) 25% ethanol, 75% buffer (iii) 50% ethanol, 50% buffer (iv) 75% ethanol, 25% buffer (v) 100% ethanol, 0% buffer

Before coming to the lab, you will need to calculate how much volume ethanol and/or potassium phosphate buffer is to be added to each test tube. Mix each solution and measure the absorbance at 625 nm. When measuring the absorbance, allow the solution to sit for one minute in the cuvette inside the spectrometer to allow undissolved material to settle and air bubbles to dissipate; both will result in a false increase in absorbance. Construct a graph of absorption at 625 nm vs. % final volume of ethanol. Thermal Denaturation—Addition of Heat Bring water to boil in a large beaker. To 10 test tubes, add 2 mL of your protein solution. Before each transfer, swirl the protein solution in order to evenly suspend any undissolved material. Each of these test tubes will be heated by placing them in the boiling water bath for the following times: 0, 5, 10, 15, 20, 25, 30, 40, 50, and 60 seconds. After heating each test tube immediately place it in a cold water bath to prevent extended heating. Measure the absorbance at 625 nm. When measuring the absorbance, allow the solution to sit for one minute in the cuvet inside the spectrometer to allow undissolved material to settle and air bubbles to dissipate; both will result in a false increase in absorbance. Construct a graph of absorption at 625 nm vs. time heated. Structural Changes Due to Denaturation For this part of the experiment, you will need a size exclusion chromatography (SEC) column, packed with Sephadex and equilibrated by the demonstrators. Allow the buffer to run out of the column to expose the upper surface of the Sephadex. Add 250 µL of your unaltered protein solution to the top of the column using a micropipette. Wash the protein solution into the column using a drop of buffer. Once the protein


(the coloured band) is completely in the Sephadex, you can add more buffer to the top of the column, being careful not to disturb the surface of the beads. Measure how long it takes for the coloured band of protein to pass completely through the column. Repeat the above experiment using a thermally denatured sample of your protein solution. If time allows, do additional tests using samples of the solution that have been denatured by other methods mentioned in this lab. Part 3: Protein Assays—The Biuret and Bradford Assays To determine the protein concentration of an unknown solution (measured in mg of protein per mL of solution) a standard curve of absorbance versus protein concentration must be constructed using protein solutions. Standards for the Biuret assay and Bradford assay will be made using stock solutions of bovine serum albumin (BSA) with concentrations of 20 and 0.5 mg per mL, respectively. Note: The 0.5 mg/mL stock solution is the 20 mg/mL stock solution after a 40-fold dilution. Note: The stock BSA solutions will have approximate concentrations of 20 and 0.5 mg per mL. Be sure to record the actual concentrations, as you will need to determine the actual concentration of each solution. Bradford Assay As above for the Biuret assay, with the following exceptions:

(i) Use the 0.5 mg/mL stock solution to prepare the standards (ii) Use Bradford reagent in place of Biuret reagent (iii) Use the 40-fold diluted protein solution of unknown concentration (iv) Allow solutions to stand for 10 minutes in place of 30 minutes (v) Measure the absorbance at 595 nm in place of 545 nm

Make dilution curve of BSA, starting with 2.0 mg/mL BSA (100%). Other points will be 75%, 50%, 25%, 5%, and 0%.


Waste Disposal Dispose of all test solutions in the appropriate waste containers in the fume hoods. Ensure that rinsing of equipment containing test solutions are also poured into the appropriate waste containers.

chemistry 3251 biology 3252 biochemistry i 2006 laboratory...

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