Life Sciences Curriculum

LSM 1101 Biochemistry of Biomolecules

2009/2010 Semester I

CONTENTS Minimal Safety Requirements…………………………………… 2 Introduction Some points on the laboratory course.....................................................................… 3 The Centrifuge............................................................................................................. 7 Pipettors...................................................................................................................... 8 Spectrophotometry……………………………………………………...............…… 11 Practical 1: pH and Buffers The Henderson-Hasselbalch equation and buffers………………………………….… 13 The pH meter………………………………………………………………………….. 14 Titration of Histidine monohydrochloride and determination of its buffering region… 15 Effect of buffer pKa on buffering capacity………………………..…………………… 17 Effect of temperature on the pH of a buffer…………………………………………… 18 Practical 2: Quantitative Protein Estimation Ultraviolet absorbance of protein…………………………………………………….. 19 Dye binding method………………………………………………………………….. 20 Practical 3: Enzyme Kinetics To plot the absorption spectra of NAD+ and NADH………………………………… 22 To determine the Km and Vmax of lactate dehydrogenase…………………………….. 23 Practical 4: DNA Gel Electrophoresis Separation of different sizes of DNA by agarosse gel electrophoresis………………. 29

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MINIMAL SAFETY REQUIREMENTS • Lab coats must be worn at all times in the lab, but removed before

leaving the laboratory • Smoking is strictly forbidden • Suitable footwear must be worn to protect against spills; do not wear

sandals or slippers

• Long hair should be tied back

• Do not eat, drink or apply cosmetics in the lab (especially nail polish – flammable solvents)

• Never use an open flame in the vicinity of flammable solvents • Never leave a lighted burner unattended

• If you have an abrasion or cut on your hands, cover it before you begin

work

• Report any accident to the demonstrators immediately • Never pipette anything by mouth • Wash your hands thoroughly with soap and water before leaving the lab

• Familiarize yourself with any additional rules relating to the laboratory in

which you are working. Note the location of eye wash points, safety shower, fire extinguisher and first aid cabinets

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INTRODUCTION SOME POINTS ON THE LABORATORY COURSE GENERAL OBJECTIVES OF THE LABORATORY COURSE Biochemistry is essentially an experimental discipline, and it is important to obtain proficiency for its techniques at an early stage if you wish to pursue the subject further. However, even if later you wish to proceed elsewhere, the skills you will learn in this course will remain useful to you in that, for the most part, they have become universally used in the other biological sciences. You will notice that each practical session has its objectives specified. It will be a matter for your judgment and comment, in conjunction with discussions with the staff, as to whether the objectives have been fulfilled. ATTITUDES TO PRACTICAL WORK It is important to approach the practicals in the right spirit, for an early feel for the way biological substances, cells and tissues are handled, will stand you in good stead. You should aim at doing as many of the manipulative procedures as possible with your own hands. To gain the maximum benefit it is important to discuss any difficulties with your demonstrator or lecturer. Try to plan your work ahead as much as possible; this means reading the text of the practical and anticipating the various steps you have to perform. The numerical result of an experiment, in isolation, is of course meaningless; the process is incomplete without interpretation, which is an aspect you should also develop. Expertise in this may come slowly, but will be guided where possible. GENERAL SAFETY IN THE LABORATORY THE GOLDEN RULE IS TO ASK SOMEBODY IN AUTHORITY ABOUT SAFETY ASPECTS IF YOU HAVE DOUBTS, AND NOT TO PROCEED UNTIL YOU ARE SURE OF THE SAFETY OF ANY PROCEDURE • Inflammable and volatile liquids (acetone, ether, hexane, benzene, alcohol, etc.) should

not be used if there is an open flame in the laboratory. • Do not heat a test tube such that it bumps (use boiling beads or a large bore tube). • Do not mouth pipette poisonous and corrosive materials. Never mouth pipette human

serum or other body fluids. • Wash away an irritant from the skin and/or eyes with copious volumes of cold water from

the sink in the first instance. Then seek advice from the demonstrator about any treatment.

• Any cuts or burns should be similarly reported. • There must be no drinking or eating in the laboratory. • Specific safety precautions will be notified or reiterated in the relevant practical sessions.

Incidentally, other potential hazards to biochemical researchers include needlestick injuries, bites from animals, infection with pathogens (viruses, bacteria) in tissues and fluids, and penetration of mucous membranes by hazardous aerosols. All are preventable with common sense precautions and a basic appreciation of laboratory safety.

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EQUIPMENT AND GLASSWARES PROVIDED • Centrifuge and centrifuge tubes • Pipettors and tips • Spectrophotometer and cuvettes • Basic preparative and volumetric glasswares • Test tubes and racks • Gel sets and power packs • Any other glasswares and equipment needed for the practical sessions

YOUR OWN SUPPLIES

• Notebook with hard covers (to resist spillages) • Paper tissues for mopping up spillages and for many miscellaneous uses • One or two cloths or rags for housekeeping • Marker pen (water-resistant) • Graph paper

SOME GENERAL POINTS OF ETIQUETTE AND GOOD LABORATORY PRACTICE • Do not crowd round another worker, both for safety reasons and to avoid distraction. • Keep any article liable to be damaged off the bench. This includes textbooks and

clothing. • Keep raucous behaviour for outside the laboratory. It is not appropriate within it. • Never pour excess reagents into the original bottles, to avoid possible contamination.

Take small quantities for your individual use. • Replace stoppers from common reagent bottles immediately. • Put everything back where you find it. • Always clean your apparatus before leaving the laboratory. If it becomes exceptionally

greasy and cannot be cleaned with detergent, ask the laboratory staff for chromic acid to effect thorough cleansing. Seek special instructions on the handling of the chromic acid.

• Wipe down the bench before you leave. • Dispose of waste appropriately. This takes some experience but briefly the following is

the pattern: 1. Corrosive wastes - into the sink with a good flow of water from the taps. 2. Organic solvents - except in small quantities, this is a specialist procedure. With

modern plastic piping, discharge of large volumes is hazardous. Pockets of vapour which are potentially explosive may also gather in drains.

3. Solids should not go into the sink, but into bins. If they are biological or sharp, see below.

4. Biological waste will be disposed of into specially marked plastic bags. Entirely liquid biological waste (such as urine) can be disposed of in the sink with copious water from the taps.

5. Sharps (broken glass, blades, needles) should go into a specially-labelled disposal box.

This is also an aspect of laboratory safety (see above).

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UNITS AND SIGNIFICANT FIGURES The scientific community insists on uniformity of units in the interests of unambiguousness and ease of interpretation. In general this course will require SI units. Because some non-SI units are inordinately useful in biochemistry their continued use is permissible. These are:

Quantity Name Symbol

time minute hour

min h

volume litre milliliter microlitre

l or L* ml or mL* μl or μL*

temperature degree Celsius °C In addition, concentration is sometimes still expressed as g/100 ml. “%” also means quantity of substance dissolved in 100 ml solution (not solvent). The concentration M for mol/l is still often used, especially in scientific journals. The formal is never used for the constituents of body fluids but can be used for lists of reagents, for the sake of brevity, as is done in this manual. But do not make up your own units. W/v and w/w are self-explanatory terms giving the relative proportions of solids and liquids in reagents. In particular never put a plural for a unit, nor a full stop, e.g. mol/l (not mol./l. or mols/l or any similar combination). As important as the correct use of units is the choice of significant figures appropriate to the accuracy and precision of the method. [Note, accuracy refers to how near the result is to the “true” result; the true result is only ascertainable by a reference method and is a concept of minimal importance in much of biochemistry, where comparisons over time and/or between groups or individuals are more important. Precision refers to the repeatability of a result; that is’ the closeness of a number of measurements on the same material.] If you report a serum glucose result as say 5.2 mmol/l you are reporting two significant figures, which are the digits required to show the precision with which a measurement can be made. To report 5.2 implies a result nearer 5.2 than 5.1 or 5.3, or more exactly, you believe the result is between 5.15 and 5.25. Were you to report, say, 5.23 mmol/l, this would imply a result between 5.225 and 5.23, which already seems optimistic for a biological assay. Zeros to the right of the decimal point are significant but those on the left may or may not be. For example in 0.003 and 0.0030 the zeros are meaningful bit in 3000 the digit 3 may be the only significant figure, implying a result somewhere between 2,500 and 3,500. This difficulty is removed by using exponents, for example 3.0 x 103 specifies two significant figures. The most skilled operator dealing with biological substances can rarely achieve a repeatability better than 3% - 5%. Therefore your choice of significant figures should be appropriate to this. This may be a difficult concept at first but at a minimum it is important not to be deceived by the capacity of electronic calculators. Lists of recommended significant figures for substances in human body fluids are published by authorities in the field and can always be consulted if need be. An example is the excerpt below.

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In print, the upper case “L” is often used to avoid confusion between one (“1”) and lower case “l”. Examples of Conversions to systéme international (SI) Units

System †

Component Present Reference Intervals

(Examples)

Present Unit Conversion Factor

SI Reference Intervals ‡

SI Unit Symbol §

Significant Digits

Suggested Minimum Increment

P Acetaminophen Therapeutic Toxic

02-0.6 >5.0

mg/dL mg/dL

66.16 66.16

13.40 >300

μmol/L μmol/L

XX XXO

1 μmol/L 10 μmol/L

B, S Acetone Negative mg/dL 172.2 Negative μmol/L XXO 10 μmol/L S Alanine

aminotransferase (ALAT)

0.35 (37°C) Units/L Karmen units/mL Reitmann-Frankel units/mL

1.00 0.482 0.482

0.35 … …

U/L U/L U/L

XX XX XX

1 U/L 1 U/L 1 U/L

S Albumin 4.0-6.0 g/dL 10.0 40.60 g/L XX 1 g/L S Aldolase 0.6 (37°C) Units/L

Sibley-Lehninger units/mL

1.00 0.7440

0.6 …

U/L U/L

XX XX

1 U/L 1 U/L

NOTEBOOK AND PRESENTATION OF RESULTS It is essential to record as you go along and not rely on memory. There is scope for creativity in all laboratory work but at this stage it lies mainly in your personal report. It should however always be headed with the title of the experiment and the date. The practical instructions should NOT be copied down in the report. Instead you should note your results, showing clearly how you worked them out, prepare charts where appropriate, and relate any difficulties or comments. You are free to make suggestions as to improvements or even to question the rationale, objectives and conduct of the session. PRESENTATION OF BIOCHEMICAL DATA Experimental data are best presented in the form of a table and/or graph. Tables and graphs must • have an informative title • be numbered with Arabic numbers, e.g. Table 1, Fig 2 • have labels for the columns and axes with the quantity and units used in the

experiments. All measurements should be in SI units. The units used should be adjusted so that the smallest number of digits is used, e.g. a concentration of 0.0045 mol l-1 is expressed as 4.5 under the heading “Conc (μmol l-1)”. If you have a wide range of values, use multipliers or exponents e.g. 5.0 x 10-2 or E-02. Refer to “Units and significant figures”.

In addition, all graphs should have • correct choice of co-ordinates. In most experiments, the change of one parameter affects

another which is measured. The known quantity is called the independent variable and the measured quantity the dependent variable. Measurement of absorbance is a common biochemical technique and the absorbance is dependent on the amount of absorbing substance. Thus the absorbance is the dependent variable.

• In all graphs, the independent variable should be plotted on the horizontal (x) axis (the abscissa) and the dependent variable on the vertical (y) axis (the ordinate).

• a suitable scale. Select a scale which will accommodate all the data points within the area of the graph paper. When extrapolation is anticipated, e.g. Lineweaver-Burk plot for analysis of enzyme kinetics, sufficient space must be left for extension and interception.

• clear, defined symbols to show each determined data point. • a smooth continuous curve (use a flexible rule) or a straight line by joining all the points

together.

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THE CENTRIFUGE The centrifuge can be used not only to separate solid particles from solutions but also non-miscible liquids of different densities. It is important to know how to use the centrifuge properly and safely. Improper use will damage it and could lead to personal injury. OPERATION 1. Before placing the tubes in the holders, make sure that the rubber cushions are placed

correctly in the tube holders. If you do not, the tubes may break. 2. It is absolutely necessary to balance the loads on a centrifuge head or rotor before

spinning. Each tube must be balanced by a tube of equal mass, diagonally opposite. Either divide your sample into an equal number of tubes or if you have only one sample or an odd number of tubes, balance the sample tube with one containing water of the same total mass.

3. When starting the centrifuge, increase the speed control gradually. Never start at high or full speed setting.

4. Never open the lid while the centrifuge is running and do not try to slow the rotor with your hand or fingers. At best this will cause re-suspension of the precipitate, at worst bruised, cut or broken fingers and an expensive repair bill.

5. If heavy vibration occurs, switch it off immediately. Do not walk away. The centrifuge may move across the bench and could fall off. Hold it firmly but gently if necessary. Do not open the lid until the centrifuge has stopped completely.

6. If a tube breaks during centrifugation, you must switch it off immediately. When the rotor has stopped spinning, every bit of broken glass and debris must be removed from the centrifuge and tube-holder. The rubber cushion must be taken out of the tube-holder and both washed and dried before reassembly.

CALCULATION OF RCF (RELATIVE CENTRIFUGAL FORCE) The force on a particle depends on the square of the velocity of rotation and its distance from the axis. It is convenient to express it relative to the force the same particle experiences under gravity. This gives the equation: RCF = (1.119 X 10-5) (rpm)2 (r) where rpm is revolutions per minute and r the radius (distance from the axis) in cm. RCF is expressed as “a number times gravity, g” e.g. 5000 x g

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PIPETTORS In clinical and research laboratories, mouth pipettes have mostly been replaced by specially designed pipettors. These pipettors are designed to measure and transfer liquids accurately and safely. They are expensive, which is why you still have to use transfer pipettes for much of your work. Currently, the Gilson or Eppendorf Pipettors, P10, P20, P200 and P1000 are provided in the biochemistry laboratory. These numbers represent the maximum volumes of respectively 10, 20, 200 and 1000 μl intended for use on each pipettor. This is clearly labeled on the push-button. Pay careful attention to the use of pipettors. You must know how to operate the pipettors correctly, as it will affect all your subsequent experiments. OPERATION 1. Ensure that you have chosen the correct volume pipettor, e.g. P10 for 1 to 10 μl, P20 for

2 to 20 μl, P200 for 20 to 200 μl, P1000 for 200 to 1000 μl. 2. Setting the volume: The pipettor consists of two or three numbers and is read from top

to bottom. The numbers indicate the volume selected, e.g. 100 is 100 μl. The volume is set by turning the adjustment ring slowly to increase or decrease the volume, making sure not to overshoot the mark.

3. Press a plastic tip firmly on the shaft of the pipettor to ensure a positive, airtight seal. 4. Aspiration: Press the push-button to the first positive stop. Holding the pipette

vertically, immerse the tip into the sample liquid. Next, release the push-button slowly and smoothly to aspirate the sample. Wipe the outside of the tip, without touching the orifice.

5. Dispensing: Place the end of the tip against the inside wall of the vessel at an angle. Press the push-button to the second stop to expel any remaining liquid. Remove the pipettor and release the push-button.

6. Eject the tip by pressing the tip ejector button. PRECAUTIONS AND CARE • Always press and release the push-button slowly and smoothly. • Never turn the pipettor upside down. • Never lay the pipettor on its side when there is liquid in the tip. • All the above precautions are to prevent liquid from entering the shaft of the pipettor.

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PIPETTE CONTROLLER

CAUTION: Always use a pipette controller whenever you are working with biological, caustic or toxic samples. Do not pipette with your mouth.

OPERATION AND PRECAUTION 1. The pipette controller serves to facilitate the pipetting of liquids with graduated pipettes,

volumetric pipettes and blow-out pipettes of either glass or plastic composition within the volume range of 0.1 ml to 100 ml. If the instrument is correctly used, the liquid comes into contact with the pipette only.

2. The pipette controller must not be used with vaporous liquids that are incompatible with polypropylene (PP), polytetrafluorethylene (PTFE) and silicone.

3. Pipetting:

Compress the suction bellows – Before attaching the pipette, squeeze the suction bellows.

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4. Attach the pipette: Hold the pipette as near to its upper end as possible, and carefully insert it into the adapter until it fits tightly. Attention: Never use force. Thin pipettes are particularly liable to break. Avoid the risk of injury! Once the pipette has been securely attached, always hold the pipette controller in a vertical position, tip down.

5. Fill the pipette:

Immerse the pipette tip into the liquid. Press the pipetting level slowly upwards. Fill the pipette so that the level of liquid is slightly above the required mark. Attention: Please take care that no liquid enters into the controller. This would impair the filtering function and reduce the suction capacity. If liquid does enter the controller, change the filter.

6. Adjust the Meniscus:

Use suitable lint-free tissue to wipe the pipette tip. Press the pipetting lever down slowly, until precise adjustment of the meniscus has been achieved. Dispense: Hold the collecting vessel in an inclined position. Place the pipette tip against the inner vessel wall. Press the pipetting lever down to dispense. In case of pipettes with a waiting time: (Imprint e.g. “Ex + 15 s”) As soon as the meniscus in the pipette comes to a standstill, start waiting time as is indicated on the pipette. Wipe the pipette tip a few millimeters upward along the wall of the vessel.

7. In case of blow-out pipettes: As soon as the meniscus in the pipette tip comes to a standstill, press the small blow-out bellows to discharge the last drop. Wipe the pipette tip a few millimeters upward along the wall of the vessel. Note: In the case of large pipettes (> 50 ml) the vacuum contained in the suction bellows is not sufficient to draw in all the liquid at once. Therefore, squeeze the suction bellows again and continue drawing up liquid. After pipetting: Hold the pipette at its extreme upper end, and gently twist and pull it out of the adapter.

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SPECTROPHOTOMETRY QUANTITATIVE LAWS OF LIGHT ABSORPTION When light passes through a solution of a substance the chance of a quantum of light being absorbed is proportional to the number of quanta entering the solution, the number of molecules absorbing substance in the light and to the chance of a quantum being absorbed if it “hits” a molecule. This proportionality explains the two empirical laws of light absorption, Beer’s and Lambert’s Laws. Together they give the Combined Law (Beer-Lambert Law):- log10(I0/I) = A = ε c l I0 is the incident light, I is the emergent or transmitted light, A is called the absorbance (alternatively extinction or optical density), ε (the Greek letter epsilon) is the molar absorption coefficient, c is the molar concentration (mol dm-3), l is the path length in cm. Note that A is the logarithm of the ratio of two quantities and it has no units. ε has the units M-1.cm-1 or mol-1.dm3.cm-1. Note that you also need to specify the wavelength, in nm, which can be done with a subscript, e.g. A405 is the absorbance at 405 nm. USE OF THE SPECTROPHOTOMETER Spectrophotometers are expensive. Treat them with care. Currently, the model of spectrophotometers available in the biochemistry laboratory is the Shimadzu UV-1200 series. Note that the basic operating procedure is the same for all spectrophotometers but each make or model of instrument has a different set of controls and sometimes the operations are done automatically by the instrument. SETTING UP A SPECTROPHOTOMETER 1. Read the instruction card provided on the use of the spectrophotometer. 2. Switch on the instrument. Check that the appropriate lamp for the wavelength you will

use is switched on. Under the Utilities menu, the light source could be selected. W for tungsten lamp only or D2 for deuterium lamp only. However, if auto-change is set between 295-364 nm, the light source will automatically change to the tungsten lamp at wavelength greater than this switching wavelength and to the D2 lamp at wavelength shorter than this set wavelength.

3. Leave the instrument for a few minutes to stabilize. 4. Zero the instrument. (a) Dark or zero transmission setting with no light reaching the detector. This may be done

automatically on some machines. (b) Place a reference cuvette in the cuvette holder in the light path. This should contain the

solvent with all the buffers and reagents except the substance you wish to measure. This is a reagent or solvent blank. Adjust the instrument to read zero absorbance or 100% transmission. You must do this whenever you change the wavelength setting.

5. Now place the solution containing the substance you wish to measure in the light path. Read the absorbance.

6. Remove the cuvettes from the spectrophotometer. Check that you have not spilled any liquid into or onto the spectrophotometer - if you have, wipe it up immediately. Ask a demonstrator if it is anything other than plain water.

7. When you have finished leave that lid of the cuvette chamber closed. If no one else is going to use the instrument switch if off.

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Practical points: Never: • Stand bottles or cuvettes on top of a spectrophotometer. • Overfill the cuvettes. • Mix different types of cuvette in one set of measurements. • Scratch the optical surfaces. • Hold the cuvette by the optical surfaces. Always: • Check that you have the correct cuvettes:

Glass or plastic for visible (320-800 nm) or near UV light. Silica for far UV (180-320 nm). Plastic UV for 220-320 nm on the Shimadzu Spectrophotometer.

• Use glass cuvettes for all organic solvents. • Remember that glass or silica cuvettes are extremely expensive, and can be easily

broken or scratched. • Check that the outside of the cuvette is clean and dry before measuring. • Wipe it gently with clean tissue. • Make sure you put the optical, polished faces of the cuvette in the light beam.

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Practical 1: pH and Buffers 1. OBJECTIVES

1.1 To familiarize yourself with the use of a pH meter. 1.2 To determine the buffering region of a buffer solution. 1.3 To understand the properties of buffers. 1.4 To carry out simple calculations using the Henderson-Hasselbalch equation.

2. INTRODUCTION

The behavior of most organic compounds is determined by their state of ionization and hence will depend to a large extent on the hydrogen ion concentration of its environment. Biochemical reactions are therefore critically dependent on pH and an understanding of the measurement and control of pH is of great importance in physiology, medicine and biochemical research. In humans, the pH of both the intracellular and extracellular compartments are maintained within narrow limits by a combination of buffers, transport of H+ or HCO-

3 across the cell membrane, control of blood CO2 and by excretion of excess acid or base. Similarly, in most biochemical experiments, the pH of the system is closely regulated by the use of appropriate buffers. The choice of the buffer is dictated both by its buffering range as well as its compatibility with the system under investigation.

The Henderson-Hasselbalch equation and buffers

Weak acids such as CH3COOH or KH2PO4 are only partially dissociated in solution.

When titrated with a strong base such as NaOH, the pH of the solution changes as a function of the amount of base added as shown in the figure below.

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At any point along the titration, the equilibrium can be defined by the equation

Ka HA H+ + A- Where Ka is the equilibrium constant and

[A-][H+] Ka = [HA]

The protons are mainly hydrated in solution and exist as H3O+, but for simplicity we shall refer to them as H+. Since by definition, pH = - log10[H+] and pKa = - log10[Ka], the above equation may be rearranged and expressed in the form of the Henderson-Hasselbalch equation

pH = pKa + log [A-] [HA]

It can be seen from this relationship between pH and pKa that when the concentration of acid [HA] and conjugate base [A-] are equal, pH = pKa. In addition, you will note that on either side of the pKa, the acid-base mixture resists change in pH upon addition of acid or base, i.e. it acts as a buffer. Buffer solutions are therefore mixtures of weak acids and their conjugate base. The pH meter

The pH meter is routinely used in biochemical laboratories for the accurate measurement of H+ concentration. The measurement relies on a glass membrane which is permeable to protons but not to other cations. An acid solution of know H+ concentration (A) is trapped within this membrane. When placed in the solution being measured (B), a potential difference will develop across the glass membrane, the magnitude of which is given by the Nernst equation

E = 1n RT [H3O+]A ZF [H3O+]B (R=gas constant, T=absolute temperature, F=Faraday and Z = valency of ion)

This potential difference is measured against a standard reference electrode which is usually incorporated into the pH electrode. The potential is directly dependent on the temperature and the instrument has to be corrected for different temperatures using the temperature control function. On some instruments, this is done automatically by a resistance thermometer in the solution (automatic temperature control). Besides this temperature-dependent variation, the electrode may also vary in efficiency giving less or even greater potential than that predicted by theory. The scale must therefore be adjusted to read correctly over the range of pH values being measured. From this you will realize that the pH meter needs to be calibrated regularly before use. This can be done using standard buffers of known pH.

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Use of the pH meter Before proceeding to use the pH meter, please consult a demonstrator.

Read carefully the instructions on the use of the pH meter which are available near each instrument. The glass bulb of the pH electrode is extremely fragile and should be handled with care at all times. Adhere to the following precautions when using the pH meter.

1. Ensure that the electrodes are always left with the bulb immersed in distilled

water or a suitable buffer. 2. Both before and after use, rinse the electrodes carefully by dipping into a

beaker of distilled water and then squirting with distilled water from a wash bottle. To prevent diluting the sample, remove excess water from the electrode by blotting lightly (but not wiping) with a tissue.

3. When taking measurements, check that the bulb of the electrode is completely immersed in the solution.

4. Remember to standardize the pH meter with the standard buffers provided as follows. Place the electrode in standard buffer pH 7 and standardize the pH meter to read exactly 7. Transfer the electrode to buffer of pH 4 or pH 10 (depending on whether you are measuring above or below pH 7) and re-standardize until the meter reads the correct pH.

3. EXPERIMENTS

3.1 Titration of Histidine monohydrochloride and determination of its buffering region

Histidine is a commonly occurring amino acid present in most proteins and enzymes. The pKa of its imidazole group ranges from 6-7 depending on the temperature, ionic strength and the nature of other functional groups present in its vicinity. At physiological pH, it can readily accept or donate protons. It plays an important role not only in the buffering action of proteins but also in the catalytic activity of many enzymes. Principle: Titration, a gradual addition of an acidic solution to a basic solution or vice versa, can be used to determine the number of acidic or basic groups in an unknown compound. A specific concentration of the compound is titrated with a known concentration of acid or base until the equivalence point has been reached. A graph can then be constructed on which the pH at regular intervals is plotted along one axis and the number of moles of added acid or base at these intervals along the other axis. Such a plot is called a titration curve and is usually sigmoid (S-shaped), with the inflection point - where the curve changes direction - corresponding to the equivalence point. From the pH at the equivalence point, the dissociation constant of the acidic or basic group can be determined. If a compound contains several different acidic or basic groups, the titration curve will show several sigmoid-shaped curves like steps and the dissociation constant of each group can be obtained from the pH at its corresponding equivalence point.

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You will prepare a 20mM solution of histidine and titrate with 0.05M NaOH. Plot a titration curve which you will use to determine the pKa and effective buffering range of histidine.

3.1.1 3.1.1 Materials

• Burette • Retort stand • Glass funnel • pH meter • Magnetic stirrer

3.1.2 Reagents

• Histidine monohydrochloride (given as dry salt, FW = 209.63)

• 0.05 M NaOH

3.1.3 Procedure 1. Design an experiment for the titration of histidine with 0.05M

NaOH. You will use 20ml of 20 mM histidine monohydrochloride for this purpose. Construct a table in which to record your data. The table must include a column for the values of mole NaOH/mole histidine.

2. Titrate at 0.5 ml increments of NaOH. The endpoint of titration is

pH 11.5

3.1.4 Data handling and questions

(a) Calculate the number of moles of histidine present in the solution.

(b) Plot a graph of pH (y-axis) against the number of moles of NaOH per mol of histidine (x-axis). The advantage of plotting the graph in this manner is that it is independent of the volume and concentrations of the solutions used.

(c) From the graph, determine the pKa values. There are two ways in which this may be done.

(i) By determining the center of symmetry i.e. the point of inflexion of the graph.

(ii) By determining the pH when 50% of the acid has been converted to the conjugate base i.e. after addition of 0.5 and 1.5 mol of NaOH per mol of histidine.

(d) At what pH does the acid-base mixture show maximal, buffering capacity? What do you think is the effective buffering range of this buffer?

(e) If you have reason to believe that the NaOH has not been accurately prepared, which of the two methods do you think will give you a more reliable estimate of the pKa? Give reasons for your answers.

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(f) Using the pKa values that you have obtained from the graph, calculate the pH of the solution after addition of (i) 5 ml and (ii) 12 ml of NaOH. Compare these values with your experimental data.

(g) The fully protonated form of histidine has the following structure:

(i) How many ionisable groups are present in histidine at the initial pH of the experiment?

(ii) Which ionisable groups are responsible for the observed pKa’s?

(h) Draw the structures of the ionic species of histidine that participate in cellular buffering.

3.2 Effect of buffer pKa on buffering capacity

Principle: Apart from using a pH meter and constructing a titration curve,

another common method to indicate the endpoint (the point of complete neutralization) of a reaction is by using visual indicators. In simple acid-base titrations a pH indicator (e.g. phenolphthalein) may be used, which turns pink when a certain pH is reached. Due to the logarithmic nature of the pH curve, the transitions are generally extremely sharp, and thus a single drop of titrant just before the endpoint can change the pH by several points - leading to an immediate colour change in the indicator.

In this experiment, we will make use of Universal Indicator - a blend of different indicators that exhibits smooth color changes over a wide range of pH values – to investigate how a buffer’s pKa determines how well it can buffer against acid or base.

3.2.1 Materials

• Test tubes • Disposable Pasteur pipettes • Universal pH indicator

3.2.2 Reagents

• 0.05 M potassium phosphate buffer, pH 7.0 (pKa 6.8) • 0.05 M Tris-HCl, pH 7.0 (pKa 8.1) • 0.05 M HCl

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• 0.05 M KOH

3.2.3 Procedure The buffer solutions provided are commonly used in the biochemistry laboratory.

Design a simple experiment to study the effect of pKa on the buffering efficiency of the buffers.

3.2.4 Questions

1. What conclusions can you draw from your experiments? 2. Which of these two buffers would you use if you were studying

the properties of a phosphatase which functions optimally at pH 7.2? Explain your choice.

3.3 Effect of temperature on the pH of a buffer

Principle: Temperature affects the pH of a buffer by altering the dissociation constants of the buffer molecules. Depending on the type of buffer, the pH either increases, decreases, or does not change when the solution is taken from room temperature to 4ºC (i.e. the temperature of a cold room, refrigerator, or ice bath). This temperature dependent behavior is most critical for protein or enzyme purification schemes in which both the pH and temperature of the procedure are critical to the isolation of an active form of the protein. 3.3.1 Materials

• Ice box • pH meter

3.3.2 Reagents

• 0.05 M potassium phosphate buffer, pH 7.0 • 0.05 M Tris-HCl, pH 7.0

3.3.3 Procedure

Measure the pH of the 0.05 M Tris-HCl and 0.05 M potassium phosphate buffer provided at room temperature and after cooling to 4°C by placing the solutions in an ice bath.

3.3.4 Questions

What effect does temperature have on the pH of the Tris-buffer and potassium phosphate buffer? Give an explanation for your observation.

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Practical 2: Quantitative Protein Estimation

1 OBJECTIVES

1.1 To record a protein spectrum and to estimate the protein content using a

direct spectrophotometric method. 1.2 To estimate the protein content using a dye-binding (“Bradford”) method.

2 INTRODUCTION

Proteins are ubiquitous components of all living tissues, whether of animal, plant or bacterial origin. They serve indispensable functions in cellular architecture, catalysis, regulation, contractile processes and immune defence. Proteins are intimately concerned with virtually all physiological events. Protein estimation is therefore very important in every investigation in biochemistry. For example, laboratory practice in protein purification often requires a rapid and sensitive method for the quantitation of protein. Presently, a variety of methods are available for the determination of protein content of a given sample. The methods employ different principles, and may be sensitive to interferences by certain salts, buffer components, and some solvents. Each method therefore has certain unique and useful characteristics as well as certain limitations. In the biochemical laboratory, the most widely used methods often employ photometric and/or colorimetric analyses as these methods are simple, rapid and have the required sensitivities.

3 EXPERIMENTS

3.1 Ultraviolet absorbance of protein

Principle: A common method of protein estimation depends on the measurement of the optical density of a protein solution. Proteins exhibit UV absorption spectra with a maximum at 280 nm, and this is due primarily to the tyrosine and tryptophan content of the protein. A pure protein can therefore be quantitated by measuring its absorbance at 280 nm in a spectrophotometer. The method is often used for a rough measurement of protein concentration because it is very quick and simple. Its outstanding advantage is that because it is non-destructive the protein solution can be recovered completely. 3.1.1 Materials

• 5 mg/ml bovine serum albumin (BSA) • Two samples of unknown concentration

3.1.2 Procedure

1. Prepare 10 ml of 1.0 mg/ml BSA. 2. Construct absorption spectra for the 1 mg/ml BSA and the two

unknowns by measuring their absorbances at 10 nm intervals over

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the range 250-360 nm (start measurement from the longer wavelength first).

3. Calculate the concentrations of all 3 samples using the respective absorbance values at 280nm.

A rough conversion is

Protein in mg/ml = E280 (absorption at 280 nm in a 1 cm cuvette)

This is strictly true only for proteins with E1% (absorption of a 1% protein solution) values of 10. As the absorption at 280 nm depends on the tyrosine and tryptophan content of the protein, each protein will have a different E1%. Tables giving the E1% of many proteins can be found in the literature. [For BSA, the value is 6.67].

3.1.3 Questions

1. Calculate the protein content of the unknown samples using E1%

values of 10.0 and 6.67. 2. What can you surmise by comparing between the absorption

spectra of the 1 mg/ml BSA and the two unknowns?

3.2 Dye binding method Principle: These methods are based on the noncovalent binding of dyes to proteins causing a colour change which may be measured in a spectrophotometer. The most widely used of these methods is that of Bradford using Coomassie Brilliant Blue G-250, now marketed commercially (Bio-Rad) as a protein assay kit. In this method, there is a shift in the absorption maximum of an acidic solution of Coomassie Brilliant Blue G-250 from 465 nm to 595 nm on binding to protein, and the assay measures the increase in absorption at 595 nm. 3.2.1 Materials

• 5 mg/ml bovine serum albumin (BSA) • Samples of unknown concentration • Dye reagent concentrate (Bio-Rad)

3.2.2 Procedure 1. The linear range of the assay for BSA is 1.2 to 10.0 μg/ml. 2. Prepare five (5) dilutions of BSA within this range using water as

diluent. 3. At the same time, also dilute the unknown samples to within this

linear range by referring to their concentrations estimated in part 1. 4. Pipette 800 μl (0.8 ml) of each standard and unknown sample

solution into a clean, dry tube. Protein solutions are normally assayed in duplicate or triplicate.

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5. For the blank or control, pipette 800 μl of water into the tube instead.

6. Add 200 μl of dye reagent to each tube and vortex. Incubate at room temperature for at least 5 minutes and not more than one hour.

7. Measure the absorbance at 595 nm, and plot a standard calibration curve of absorbance against protein (BSA).

8. Estimate the protein content of the unknown samples from their absorbances at 595 nm using the calibration curve.

3.2.3 Questions

1. Construct a protein standard curve based on the dye-binding (“Bradford”) method, and establish the protein content of the unknown solutions provided.

3. Compare the protein content of the unknowns using the two methods.

4. Calculate the molar extinction coefficient of BSA if a 1.0 mg/ml solution in buffer has an absorbance of 0.667, and the molecular weight of BSA is 66,000.

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Practical 3: Enzyme Kinetics 1. OBJECTIVES

1.1 To plot the absorption spectra of NAD+ and NADH. 1.2 To understand the rationale for measuring reactions of NAD+-linked

dehydrogenases at 340nm. 1.3 To examine the relationship between A340 and concentration of NADH and to

determine the molar absorption coefficient of NADH. 1.4 To understand how initial velocity of an enzyme-catalysed reaction is

obtained. 1.5 To plot the Michaelis-Menten curve and Lineweaver-Burk plot with the data

obtained. 1.6 To calculate the values of Km and Vmax of LDH.

2. INTRODUCTION

To obtain Michaelis-Menten kinetic data for an enzyme, it is necessary to control all variables except the substrate concentration. When the initial velocity is plotted against the substrate concentration, one will have the Michaelis-Menten curve, from which, however, a determination of Km and Vmax is difficult. Therefore, Lineweaver-Burk plot is often used to obtain these Michaelis-Menten kinetic data. In addition, this plot can also be used to characterize and measure the effects of inhibitors of an enzyme. Lactate dehydrogenase (LDH) will be used in this practical to illustrate how such kinetic data are obtained. LDH catalyses the following reaction:

Lactate + NAD+ pyruvate + NADH + H+

Many NAD+- and NADP-linked dehydrogenases can be measured spectrophotometrically by monitoring the absorbance of NADH or NADPH formed. The activities of other enzymes may also be determined in a similar way if the products of the catalysed reactions can be coupled to those catalysed by dehydrogenases. NADH will be used in this practical to illustrate the application of spectrophotometry for studying enzyme kinetics.

3. EXPERIMENTS

3.1 To plot the absorption spectra of NAD+ and NADH

3.1.1 Materials • 0.1 mM solution of NAD+, pH 7.5 • 0.1 mM NADH, pH 9.0.

3.1.2 Procedure

Record the absorbance of each solution between 240 to 400 nm at intervals of 20 nm. With every change in wavelength setting, zero the

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instrument with a blank containing buffer alone. Why is it necessary to zero the instrument?

3.1.3 Report and Questions

1. Plot a graph to show the relationship between absorbance (y-axis) against wavelength (x-axis).

2. What is the main difference in the absorption spectra between NAD+ and NADH?

3. What is the wavelength of choice for measuring NADH in a mixture containing both NAD+ and NADH? Give your reasons for your answer.

3.2 To study the relationship between absorbance and concentration of NADH

3.2.1 Materials

• 1 mM NADH in 0.05 M sodium pyrophosphate buffer, pH 9.0 • 0.05 M sodium pyrophosphate buffer, pH 9.0

3.2.2 Procedure Prepare a series of tubes containing final concentrations of 0.01 mM to 0.2 mM NADH. As the cuvette can accommodate 3 ml, the final volume of each diluted solution should be at least 3 ml. Read the absorbance of NADH for each concentration at A340.

3.2.3 Report 1 Tabulate your data in the form of a table. 2 Plot a graph to show the relationship between absorbance and

concentration of NADH. 3 Calculate the molar (or millimolar) absorption or extinction

coefficient of NADH, applying the Beer-Lambert equation.

3.3 To determine the Km and Vmax of lactate dehydrogenase

3.3.1 Materials • 200 mM sodium lactate • 10 mM NAD+ • 2 unit/ml LDH • 0.05 M sodium pyrophosphate buffer, pH 9.0

3.3.2 Procedure

Pipette 0.1 ml NAD+ and 30 μl of sodium lactate into a cuvette and topped it up with buffer to a final volume of 2.8 ml. Zero the spectrophotometer. Start the reaction by adding 0.2 ml of LDH. Cover the cuvette with a small piece of parafilm and mix rapidly and thoroughly by inversion. Monitor the progress of the reaction by recording A340 at 30 s intervals for 4 min. Repeat the above assay with different volumes (up to 300 μl) of sodium lactate, keeping other

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conditions unchanged. You should have at least 5 different concentrations for the substrate (from 30 μl to 300 μl).

3.3.3 Report

1 Plot the increasing absorbance against the reaction time for each substrate concentration. Find the initial velocity from the curves obtained (note: that the value obtained here will be in absorbance units per min)

2 By using the information from 3.2.3, convert the initial velocity to moles per min.

3 Plot the Michaelis-Menten curve and Lineweaver-Burk plot (note: in both plots, the initial velocity should be in moles per min). Compare and comment on the Km and Vmax values obtained from these two procedures.

Practical 3: Enzyme Kinetics 3.1 To plot the absorption spectra of NAD+ and NADH

Wavelength NAD+ NADH 400 380 360 340 320 300 280 260 240

3.2 To study the relationship between absorbance and concentration of NADH

mM NADH A340

3.3 To determine the Km and Vmax of lactate dehydrogenase (Total vol = 3 ml)

Tube No.

10 mM NAD+ (ml)

200 mM lactate (ml)

0.05 M buffer pH

9.0 (ml)

LDH (ml)

mM lactate (final conc)

1 0.1 0.2 2 0.1 0.2 3 0.1 0.2 4 0.1 0.2 5 0.1 0.2

A340 Time

(sec) Tube 1 Tube 2 Tube 3 Tube 4 Tube 5 0 30 60 90 120 150 180 210 240

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Supplementary Information for

Practical 3: Enzyme Kinetics Coenzymes: NAD+: Nicotinamide adenine dinucleotide, oxidized form NADH: Nicotinamide adenine dinucleotide, reduced form NADPH: Nicotinamide adenine dinucleotide phosphate, reduced form (NADP+: Nicotinamide adenine dinucleotide phosphate, oxidized form )

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An illustration of how NADPH is utilised in metabolism. The free energy released during metabolism is conserved by the synthesis of ATP from ADP and phosphate or by the reduction of the coenzyme NADP+ to NADPH. ATP and NADPH are the major free energy sources for anabolic pathways.

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Practical 4: DNA Gel Electrophoresis 1. OBJECTIVES

1.1 To resolve different sizes of DNA by agarose gel electrophoresis. 1.2 To appreciate the use of an enzyme (a protein) to cleave DNA.

2. INTRODUCTION Agarose gel electrophoresis is a commonly used method to separate nucleic acids of

different sizes. The separated nucleic acid fragments can then be purified and further used in molecular biological manipulations. Nucleic acids have net negatively charged phosphates with pKa of ~ 2 and hence, in neutral conditions (pH~7), the molecules will have an overall negative charge. To a great extent the molecule possess a constant charge: size ratio. Hence, nucleic acids of all sizes will migrate at a similar rate in liquids and it is not possible to separate them in liquid. With molecules possessing constant charge: size ratios, one way to separate the various sizes is by exploiting the frictional properties of the electrophoretic medium. The most commonly used method is to carry out nucleic acid electrophoresis in gels that act as molecular sieves. Gels are semi-solid matrices entrapping a buffer.

AGAROSE Agarose is a natural polymer extracted from red seaweeds. Agarobiose is the idealized subunit of the agarose molecule.

1,4 linkage

1 4

Agarose is actually a polymer of the above structure with ill-defined chemical substitutions (e.g., methyl groups on R). It is a polysaccharide of 1,3-linked β-D-galactopyranose and 1,4-linked 3,6-anhydro-α -L-galactopyranose. This basic unit is repeated to form long chains with an average MW of 120 kd, representing ~ 400 agarobiose units. There are also charged groups on the polysaccharide (notably, pyruvate and sulfates). Two commonly used buffers are Tris-acetate-EDTA (TAE) and Tris-borate-EDTA (TBE).

O OCH2OH

OH

OHO O

CH2

O

OR

3,6 anhydro linkage

n

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To detect DNA in agarose gel electrophoresis, there are a few choices. It is usual to add small amounts of ethidium bromide (< 1 μg/ml of gel) into the gel. Alternatively, after the run, the gel is stained with ethidium bromide for a short period of time ~ 1/2 hr, followed by destaining in water. What is not frequently carried out (but practised by some) is that to add ethidium bromide into the running buffer. This of course will require a large amount of ethidium bromide and spillage of the running buffer (which does happen) will contaminate the surrounding surfaces. The higher the % of gel used, the longer the time needed to resolve the fragments. It is tempting at times to run the gel at high voltage. This will invariably cause overheating (Joule heating). To estimate the sizes of the DNA fragments after cleavage by an enzyme (also known as ‘restriction digest’), known molecular weight markers are electrophoresed in the same gel.

3. EXPERIMENT

3.1 Separation of different sizes of DNA by agarose gel electrophoresis

3.1.1 Materials • Agarose gel • Gel electrophoresis equipment • λ DNA • Hind III enzyme • EcoRI enzyme • 1 Kb DNA marker (1000 base pair) • 10x enzyme buffer. • Gel loading dye.

3.1.2 Procedure

Preparation of agarose gel

1. You will be supplied with a bottle containing 1% molten agarose (w/v) and ethidium bromide (<1 μg/ml of gel) in TAE buffer. This bottle is kept at 60°C until you are ready to pour the gel. (The agarose solution was prepared by adding 1g of solid agarose in 100 ml TAE buffer and subsequently heated to dissolve the agarose). Note: ethidium bromide is a carcinogen and MUST be handled carefully! Wear gloves.

2. Assemble the gel tray by taping both ends of the tray and carefully

place the comb provided at one end. 3. Remove the molten agarose from the water-bath and carefully pour

the gel into the tray, avoiding any bubbles. If there are bubbles on the gel, carefully remove them by using a pipette tip. Reposition the comb if it has been disturbed during the pouring of the gel.

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4. Allow the gel to solidify (turning into an opaque piece of gel). Do not move the gel. In the mean time, set up enzymatic cleavage of DNA as described below.

Enzymatic digestion

You will be digesting bacteriophage λ DNA with two endonucleases, Hind III and EcoRI . Single and double digestions will be carried out. After digestion, the resulting fragments of λ DNA will be separated on the agarose gel which you have prepared above. To estimate the sizes of the different fragments, a 1 kb DNA marker (every band is 1000 base pair in length) will be co-electrophoresed.

1. Set up the following reactions in microfuge tubes (Table 1).

2. Cap the tubes and mix the contents briefly (vortex briefly). Spin down the droplets from the sides of the tubes using the microcentrifuge (~ 10 sec).

3. Incubate all tubes at 37°C for at least 30 min. Table 1: Restriction of λ DNA Reagents Tube 1

(Uncut) Tube 2 (EcoRI)

Tube 3 (HindIII)

Tube 4 (EcoRI + HindIII)

λ DNA 20 μl 20 μl 20 μl 20 μl 10X Buffer 4 μl 4 μl 4 μl 4 μl Sterile Water 16 μl 13 μl 13 μl 10 μl Restriction enzyme 0 μl 3 μl 3 μl (3 + 3) μl

Gel electrophoresis

At the end of the digestion, the λ DNA fragments can then be separated on the agarose gel.

1. Untape the agarose gel and carefully remove the comb. Note that

wells are formed when the comb is removed. Place the gel in the gel tank. Ensure that the orientation of the gel with respect to the polarity of the electrical leads is correctly placed.

2 Fill the tank with TAE buffer till the gel is submerged. 3 Briefly spin down the contents of all tubes. 4 Add 10 μl of gel loading dye (consisting of 25% glycerol, 0.2%

bromophenol blue, 0.2% xylene cyanol and TE buffer) to each of the tubes.

5 Mix carefully and pipette 20 μl of the contents and 6 μl DNA marker into the respective wells.

6 Record the loading sequence of the samples. 7 Once the samples have been loaded, place the lid on the gel tank

such that the samples migrate from the cathode (black) to the anode (red).

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8 Run at constant voltage of 120V until the bromophenol blue band migrates at least half way down the gel. (Note the bubbles generated during electrophoresis.)

9 Stop the electrophoresis by switching off the power supply. 10 Carefully remove the gel and view under UV.

Note: consult your demonstrator.

4 Report and Questions

1. Plot a standard curve using 1 kb DNA markers. 2. Calculate the sizes of the digested λ DNA. 3. Comment on the advantages and disadvantages of using (a) Agarose (b)

Polyacrylamide gels for the separation of DNA molecules. 4. Why is the digestion of DNA with enzymes Hind III and Eco RI also known

as “restriction digest”?