University of Northern British Columbia
Biochemistry Laboratory
UNBC Biochemistry Laboratory
Isolation and Purification of myoglobin from ground beef
(Labs 5, 6, 7, 8, and 9)
This lab will run over a period of 5 weeks. At the end of 5 weeks you will write a
scientific manuscript on your findings.
Myoglobin is a member of a family of proteins collectively called globins. Along
with hemoglobin, it plays a vital role in the acquisition and utilization of oxygen in
animals. These vital proteins can deliver oxygen efficiently to cells and store it for later
use for the oxidation of foodstuffs. Myoglobin is the oxygen storage protein, and
hemoglobin is used for the oxygen transport. Their relationship gives some important
insights into how protein function evolves, as we know more about the structure, function
and evolution of the globin family than about any other group of proteins. The structures
of myoglobin and hemoglobin were amongst the first protein structures to be determined
by X-ray diffraction. The ability to determine the tertiary structure of proteins was a
major breakthrough, as scientists seek to understand the functions of proteins by studying
their structures and establishing their essential chemical and structural features that lead
to efficient function.
There is a common fold to these proteins, with hemoglobin being a tetramer of
myoglobin-like chains. A single polypeptide chain is folded about a prosthetic group, the
heme, which contains the oxygen binding site. The function of these proteins depends on
the chemical equilibrium of oxygen binding to the iron ion that is chelated to this heme
group. Myoglobin is a monomeric polypeptide of 153 amino acid residues. Eight helical segments form the tertiary structure of the protein, a nearly spherical shape with a
diameter of ~30A. The main function of the polypeptide is to stabilize the bound heme
group for oxygen transport. Myoglobin is present in large concentrations in muscles,
particularly in those in which oxygen deficiency could be life threatening. The human
heart contains about 0.1% myoglobin by weight. The high concentration of myoglobin is
responsible for the color of mammalian red muscle tissue, i.e. the red colour of
hamburger. This high concentration also provides a good source for purification.
Learning Outcomes
o To learn how to make buffer solutions.
o To isolate and characterize myoglobin from ground beef
o To develop expertise and an understanding of the techniques utilized in Labs 7, 8
and 9.
o To write a formal lab report in the form of a journal article
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Biochemistry Laboratory
Lab 5: Making buffers and pH, using a pH meter and
preparing a crude extract from ground beef
Introduction
A buffer is a solution that resists changes in pH when a small amount of acid (H+)
or base (OH-) is added. A typical buffer system consists of a weak acid (proton donor)
and its conjugate base (proton acceptor) sometimes referred to as a conjugate base-pair.
Buffers are extremely important in biological systems as they have a major role in
stabilizing the pH of living systems. Many biochemical processes produce or consume H+
ions and a good buffer system controls the biochemical environment. An artificial buffer
system is found to be the best substitute for a natural cell environment.
The expression of acidity or alkalinity is in pH units, where pH is defined as the
negative log of the hydrogen ion concentration, pH = - log [H+]. This is a convenient
logarithmic way to express the hydrogen ion concentration in a solution as it eliminates
dealing with large powers of 10 and compresses the range of concentrations onto a scale
between 1 and 14. Solutions having a pH greater than 7 are alkaline (basic). Solutions
having a pH of less than 7 are acidic. At pH of 7, the solution is considered neutral.
There are two common ways to determine pH. The first is to use pH indicators,
dyes whose color is pH dependent. They can be in solution, or impregnated into strips of
paper called litmus strips. As the dye color will depend on a bell curve, indicators are
used only for approximation of pH. The second method is using a pH meter. A pH meter
is a potentiometer or voltmeter which will measure the H+ concentration in solution. It
measures the potential difference between a reference electrode and a sensory electrode.
Today a combination electrode has incorporated both functionalities with a single
electrode. There is a glass membrane which acts as if it is selectively permeable to H+
while other cations and anions are excluded. This selectivity results in a potential across
the membrane that is a linear function of pH, and can be measured.
The strength of an acid is determined by how much of the hydrogen ion
dissociates when the acid is dissolved in water. If HA is a weak acid, then it
disassociates as:
HA
H+ + A(Rx'n.1)
+
Where HA = undissociated acid, H = hydrogen ion, and A = conjugate base
Thus, the equilibrium constant (Ka) is as follows:
k
H A
(Eq.2)
Ka  1 
HA 
k2
The larger the Ka , the greater the dissociation and the stronger the acid. Since the [H+]
concentration can be expressed as pH (pH = -log [H+]), the Ka can be expressed by an
equivalent expression pKa= -log Ka. Rearranging Eq.2 to solve for [H+],
K HA 
H  a 
(Eq.3)
A
  
 
 
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When converted to log form,
 
log H   log K a  log
Biochemistry Laboratory
A 
-
HA 
(Eq. 4)
With our prior expressions, Eq 4 can be reduced to :
A(Eq. 5)
pH  pK a  log
HA 
This is known as the Henderson-Hasselbach equation which correlates pH and pKa and is
a mathematical equation for a weak acid behavior. When [A-] = [HA], then
HA   0 .
log
Aand pH = pKa. This is the pH where there is an equal number of positive and negative
ions. It is the equilibrium constant derived from the rates of association and
disassociation of a weak acid or (base), expressed in log form. As pKa is the pH at which
the buffer experiences little change in pH
upon addition of acids or bases, it is the
12
center of the buffering system and is the pH
at which the buffer is most effective. The
1
0
most effective pH range for a buffer is pKa
8
±1. The pKa value also indicates the relative
pH6
strengths of an acid, the lower the pKa, the
4
greater the relative strength (See Fig 5.1).
2
The Henderson-Hasselbach equation
allows us to calculate the pH of a buffer
-0.6
-0.4
-0.2
0.0
0.2
NaO
system or the pKa of a weak acid or base in
HCl
mmoles of titrant
H
preparation of a buffer. Generally, when we
added
Fig 5.1. Sample titration curve of a buffer
are making buffers, we know the pH needed,
we find the pKa of the weak acid, and use the
Henderson-Hasselbach equation to determine the quantities of acid and base needed.
The pKa values for any particular molecule is generally determined by titration.
The free acid of the material is titrated with a suitable base and the titration curve is
recorded. The pH of the solution is monitored with a pH meter as increasing quantities of
base are added. If the moles of titrant are plotted vs the pH, there will be a sigmoid curve,
where the point of inflexion indicates the pKa value (Fig. 5.1).
Buffer capacity refers to a measure of how well a given buffer protects against
change in pH. It depends largely on the concentration of the buffer solution. The
maximum buffering capacity of any buffer is always found at pH = pKa, and usually
defined as the number of equivalents of either H+ or OH- required to change the pH if a
given volume of buffer by one pH unit.
 
 
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0.4
University of Northern British Columbia
Some criteria in choosing a buffer system:
• Choose a buffer with the appropriate pKa (
Table 5.1). The closer the pKa of the buffer
system to the pH of the working environment
the greater the buffering capacity of the
buffer. However a pKa between the 6.0 to
8.0 region is useful for most biological
reactions
• High aqueous solubility
• Minimal penetration of biological
membranes
• Minimal salt effects or effects due to the
ionic composition of the solution
• Minimal effects due to concentration or
temperature
• Minimal interactions with mineral cations
• Enzymatic and hydrolytic stability
• Minimal absorbance between 240 and 700nm
• Easy to purify
• Non-toxic
• Minimal participation in biochemical
reactions
Biochemistry Laboratory
Table 5.1 Common buffers and
their pKa values*
Compound
pKa value
Oxalic acid
1.27
Histidine
1.82
Phosphoric acid
2.15
Glycine
2.35
Citric acid
3.13
Citric acid
4.76
Acetic acid
4.76
Histidine
6.04
MES
6.09
Citric acid
6.40
Phosphoric acid
6.82
MOPS
7.15
TES
7.40
HEPES
7.47
Tris
8.08
Bicine
8.26
Pyrophosphate
9.95
CHES
9.50
Glycine
9.78
* Taken from Ninfa and Ballou, 2004
There are two ways to prepare a buffer. One method is to titrate a weak acid with
a strong base, or a weak base with a strong acid. In this procedure, one would make the
solution slightly more concentrated than needed, add the acid or base until the pH is
correct, and then dilute the solution with the appropriate amount of solute to the correct
concentration. The Henderson-Hasselbach equation is used when there is no pH meter
available or the biochemical system will not allow the use of one. This would require
known amounts of the A- and HA forms to be mixed and diluted to volume, and no
titration is required.
Before using the pH meter, read the instructions (Appendix B ). Before starting with the
lab, turn on the pH meter and standardize it using the standards provided. A few other
notes in pH meter use are as follows:
1. Always check the liquid level in the electrode before use. It should be slightly
below the fill hole.
2. Always rinse the electrodes with deionized water and blot dry with a KimWipe
before use and every time a new solution is measured.
3. Standardize the meter with at least 2 standard buffers, one with pH below and one
with pH above the values to be measured.
4. The bulb of the electrode must be completely covered with the solution.
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5. Do not allow the electrode to remain in any solution, especially proteins, any
longer than necessary.
6. Rinse the electrode with deionzied water immediately when finished, and return
electrode to the storage solution if not going to be used for short time periods. If
finished with the meter, switch to “off” or “standby”.
7. There are many types of electrodes with a wide variety of applications. Choosing
the right type is critical when there are certain restrictions such as high
temperatures, high pH or salt concentrations, high or low ionic strength, Clinterference or emulsions.
Lab Safety
PPE: Wear lab coat, goggles, and gloves
Chemical hazard: Low, you will use Tris, KH2PO4
Biological hazard: None
Physical Hazard: None
Exercise 5.1: Making a buffer for the extraction of myoglobin
in ground beef
You will need two buffers for the extraction of myoglobin from ground beef. The first
buffer will be 20 mM, pH 5.6, KH2PO4 (MW= 136.09 g/mol) (Buffer A) and the second
buffer 20mM Tris buffer pH 7.5 (MW= 121.1 g/mol) (Buffer B). However, due to time
constraints two groups will collaborate to make these buffers. Therefore each group will
make one of the two buffers and will subsequently share their buffer solutions for labs 6,
7, and 8. But make sure you do calculations for all buffers and solutions.
1. Weigh _____g to make a 50mL 0.5M ________solution. Show the calculations for
both buffers in your lab notebook.
Sometimes you need to make a buffer using a very small amount of solid
chemicals. This amount can be so small that you cannot weigh it
accurately. To solve this problem you can make a solution of a much
higher concentration and then use a small amount of the concentrated
solution for your buffer.
2. Take _____mL of the 0.5M solution you made in step 1 and add 200mL deionized
water (ddH2O/ MilliQ) to a 400mL beaker. Show the calculation in your lab
notebook.
3. After calibrating the pH meter insert the pH probe into the solution so that the bulb is
submerged.
4. Make sure all the chemicals are completely dissolved before you start to add base or
acid to adjust the pH.
5. With the pH probe in the buffer solution add either 0.5 N base (KOH) or acid (HCl)
6. Once the desired pH is reached transfer the solution to a 250mL volumetric flask and
adjust the volume with ddH2O to 250mL. Label this flask.
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Proper labeling will include information on the buffering compound, its
concentration, pH of buffer solution, date, and initials of the person
who made it.
Exercise 5.2: Isolation of myoglobin from ground beef muscle
1.
2.
3.
4.
5.
Weigh 10 g previously frozen, thawed ground beef into a 50mL conical tube.
Mix with 20.0 ml, 20 mM, pH 5.6, KH2PO4 (Buffer A).
Centrifuge at 20 ºC for 20 min at 5000 rpm in large centrifuge tubes.
Save 1.0 ml of supernatant on ice in a microfuge tube (Crude extract 1).
Filter supernatant through glass wool using a funnel and beaker. Discard pellet in
appropriate container.
6. Filter filtrate through a 0.45 m syringe filter into a 15.0 ml cylinder. You may
require more than one syringe filter.
7. Record the total volume to nearest 0.1 ml.
This is so that you can determine the total amount of protein
8. Aliquot 1 mL to 1.5 microfuge (Filtrate 2), 1.8 mL to a 2mL microfuge tube (will be
used in lab 6). The remainder you will use in Exercise 5.3.
Exercise 5.3: Acid digestion of filtered extract to be used
for Iron analyses
**The following procedure should be done in a fume hood.**
1. Transfer 10 mL of the filtered extract to a 100mL glass beaker and add to this 5 mL
6N HNO3.
2. Place a watch glass over the beaker and start to heat the extract very gently. It is
important to heat the sample to a gentle boil and not a vigorous boil.
You will notice that the colour will change over time and that the solids
will dissolve. The solution will go from a dark brown to light brown and
eventually a light yellow.
3. Once the solids are dissolved remove the watch glass and keep boiling (gently) until
the volume is less than 5mL.
4. Quantitatively transfer the solution to a 5 mL volumetric flask while rinsing with
deionized water and add ddH2O up to the 5 mL mark.
5. Transfer this concentrated solution to a clean 15 mL conical tube and label it with
your name, date and type of sample.
6. You should have four samples in total: Sample 1 and 2 in 1.5 mL microfuge tubes, a
2mL sample in a 2mL microfuge tube and then 5 mL of acid digested extract in a
15mL conical tube. Give these samples to the instructor for next week.
**Remember to give your buffers to the instructor for safe keeping.**
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Biochemistry Laboratory
Lab 6: Cation exchange chromatography of ground beef
extract
Learning Outcomes
In this experiment, you will be purifying the protein myoglobin from beef muscle
using ion-exchange chromatography.
Introduction
In this experiment you will be isolating the protein myoglobin from bovine
muscle, purifying it and identifying it using the techniques of ion exchange and
electrophoresis. You will be using an ion exchange column to extract the protein
myoglobin from ground beef. This column is packed with a resin called Sepharose CM
(carboxymethyl). This resin is a weak cation ion exchanger which carries a negative
charge above the pH of 4.5. It has an affinity for molecules with a positive charge.
Specific ions are eluted (removed) from the column by either changing the pH or
changing the ionic strength of the eluent.
Ion exchange chromatography is a second chromatographic technique for
separating out biomolecules (bioseparation technique). In ion-exchange, molecules are
separated on the basis of their electrical charge; as oppositely charged particles are
attracted to one another. Ion exchange separations, as for other chromatographic
techniques, take place in columns packed with an ion-exchange resin. These are resins
which are either polyanions (possessing positively charged ions) or polycations (
possessing negatively charged ions). The resins consist of a polystyrene or cellulose
matrix with the attached functional group. The most important part of the ion exchanger
is the charged group that is chemically bonded to the support.
When selecting an appropriate ion exchange resin, deciding on whether to use a
cation or anion exchanger is dependent on what is to be purified, the pH necessary to use
in the column, and if the functional group is to be strong or weak. A strongly acidic or
cationic exchanger would be sulfonic acid, and a strongly basic or anionic exchanger
would be a quaternary amine. Amino acids require strong ion exchangers which would
denature proteins.
In the separation of biologically important macromolecules, such as enzymes and
proteins, cellulose and cross-linked dextran (Sephadex) is the most common solid
support. To this support, charged groups such as diethylaminoethyl (DEAE) or
carboxymethyl (CM) are chemically linked to them to make anion and cation exchangers,
respectively. DEAE or CM are weak ion-exchangers, and require activation by treatment
with a buffer of suitable pH to charge the exchangers so that they have the appropriate
charge to interact with macromolecules in solution. The compounds being separated
likewise must be charged by reacting specific groups at some designated pH before the
mixture is loaded onto the column.
Proteins can be separated and purified using ion-exchange chromatography to
take advantage of their different properties under a controlled environment. Proteins
differ from one another in the proportion of the charged amino acids that they contain.
Hence, proteins will differ in net charge at a particular pH. This difference is exploited in
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ion-exchange chromatography, one of the most widely used procedures for purifying
proteins. The procedure is dependant on the formation of electrostatic linkages between
the resin and the substance being separated. This technique is particularly useful for
separating compounds of similar structure such as proteins.
If a protein has a net negative charge at pH 7, it will bind to a column containing
positively charged beads, whereas a protein with no charge or a positive charge will not
bind. The negatively charged proteins bound to such a column can then be eluted by
washing the column with a solution of NaCl at the appropriate pH, where the Cl- ions
compete with the protein for the positively charged groups on the column. Proteins
having a low density of negative charge elute first, followed by those with a higher
density of negative charge, so that individual bound proteins can be eluted at different
salt concentrations. The pH of the buffer may also be adjusted to alter the ionization state
of the amino acid side chains and hence the net charge on the protein.
After the mixture has been loaded, the desired compound is recovered by washing
the sample off the column, sometimes with buffers of varying strengths. To separate a
protein effectively, it is necessary to take into account the size of the sample, which will
determine the dimensions of the column and the volume of the eluent needed.
For this experiment, myoglobin will be separated by cation-exchange
chromatography. As the sample passes through the column, fractions of the eluent are
collected at specific times or volumes. You will determine the extract purity to identify
which test tubes have your compound of interest, and from this, you will know which
samples to use to analyze in the next lab with electrophoresis.
Lab Safety
PPE: Wear lab coat, goggles, and gloves
Chemical hazard: Low, you will use Tris, KH2PO4
Biological hazard: None
Physical Hazard: None
Solutions required today
o 20 mM, pH 5.6, KH2PO4 (Buffer A)
o 20mM Tris buffer pH 7.5 (Buffer B)
o MilliQ
Location
You made last lab
You made last lab
Back bench of lab
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Exercise 6.1: Ion exchange column chromatography of beef
extract
1. Preparing the peristaltic pump:
a. To prime the pump place the inlet tube into the mobile phase, switch the
pump on by pressing “S1” to the forward position.
b. Press “S2” to prime position and let the peristaltic pump run until mobile
phase appears at the end of the outlet tube. The pump is now primed.
2. To attach the peristaltic pump to the column switch the pump off first. Attach the
inlet tube to the bottom of the column. But make sure you removed the top cap
and then the spout cap from the column. Do not misplace these since you will
need them at the end of the lab.
3. To determine the flow rate of your column:
c. Place the outlet in a 10 mL graduated cylinder, turn “C1” on the pump to 5
and “S2” to slow.
4. Determine the time it takes for 8 mL of the mobile phase (eluent) to elute.
Determine the flow rate in milliliters per minute.
It is important not to let the column run dry at any time. If the
column runs dry air pockets will be created between and inside the
beads. This will lead to the formation of fissures and cracks. This will
slow down the flow rate of the column and reduce resolution. The only
remedy is to repack the column. There it is important to add mobile
phase to the column to prevent it from drying out. When adding mobile
phase make sure to add it slowly so that you do not disturb the column
bed.
5. Equilibrate the ion exchange column with an additional 10 ml of 20 mM, pH 5.6,
potassium phosphate monobasic (Buffer A).
6. Collect four 2 ml fractions, adding 9-10 ml Buffer A as needed and collecting 2
ml fractions in previously labeled small test tubes.
7. Check and record the absorbance at 280 and 417 nm of these fractions, using
Buffer A as a blank.
Only use 500uL of your fraction and 500uL of the appropriate
buffer to measure the absorbance. The absorption readings
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should be close to zero and should be stable (ie. should not
changes much between fractions). When this is achieved the
baseline for the chromatogram has been established.
8. Pipet 2.0 ml of crude filtered beef solution from Ex. 5.2 onto the column and
allow it to enter the CMC matrix. Start to collect 2 mL fractions
9. Immediately pipette 2 ml of Buffer A onto the column. Allow to enter column.
10. Collect 2 ml fractions, adding 9-10 ml Buffer A as needed and collecting 2 ml
fractions in previously labeled small test tubes.
11. Check and record the absorbance at 280 and 417 nm of these fractions, using
Buffer A as a blank. Note the fraction with the highest absorbance.
Only use 500uL of your fraction and 500uL of the appropriate
buffer to measure the absorbance. Keep all fractions with
absorbance readings greater than 0.1 at 280nm. You will decide
later which fraction to keep
12. Continue to collect fractions until the absorbance at 280nm is approximately zero
or stays consistently below 0.1.
13. Once A280nm = 0, remove all of Buffer A from the top of the column.
14. Immediately add 10 ml of 20mM Tris buffer pH 7.5 (Buffer B) to the top of the
column.
15. Collect 2 ml fractions as before in labeled test tubes.
16. Read and record the absorbance of these fractions at 280 nm and 417 nm using
Buffer B as the blank.
17. Continue to collect fractions until A280 nm is approximately zero.
18. Save 2 ml of each sample fraction in 2mL microfuge tubes for later use in labs 7
and 8. You need to keep a total of two fractions from the column
chromatography.
These 2 fractions should consist of one fraction from the
washing step with Buffer A and one fraction during the elution
step with buffer B. The fraction from the elution step should
be a combination of the highest 417nm/280nm ratio and the
highest absorbance at 280nm.
Even though you may discard some of the samples you will need
all the data.
19. Label all microtubes with name, date and fraction number. With the two samples
of exercise 5.1 you should have 4 samples in total.
20. Freeze for Lab #7, during which you will determine the protein concentrations of
all four fractions.
.
** Also perform Exercise 7.1**
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Lab clean up
1. Wash all glassware and test tubes and let dry and place back on cart or back into your
locker
2. Discard all liquid waste into the aqueous waste container located in the fume hood
3. Refill tip boxes and place back on cart
Lab 6: Short Lab Report
Your report will be evaluated for overall quality of the writing, grammar, spelling, the
ability to construct tables and figures (graphs) correctly and your ability to analyze
and interpret data appropriately.
For your report include the following sections (marks):
Report (18) + lab demeanor (2) = 20
o Introduction (8)
o Results (8)
o Chromatogram for the purification of myoglobin
o Appendix (2)
o Any sample calculations
Remember to write out every step of the calculations

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Lab 7: Determination of protein concentration by
Spectroscopy
Learning Outcomes
To perform two different spectrophotometric assays for the determination of
protein concentration. The first assay is a direct assay, where the concentration of the
myoglobin is determined from the absorbance and ε value at a UV wavelength of 417nm.
The second assay is colorimetric where a visible change in colour is measured as the
concentration of the protein increases. This is an indirect assay which is dependent on the
reaction between the protein and a dye. It is used for determining the total protein
concentration of a mixture of proteins.
Introduction
Spectrophotometry is the study of the interaction of electromagnetic radiation
with molecules, atoms and ions. It has become extremely important in any biochemistry
sector, as measurements of emission or absorption of electromagnetic radiation can
provide information on molecular structure. This means that spectroscopic methods can
provide insights into the physical and chemical properties of biomolecules. Qualitatively,
spectrophotometry can be used to identify biomolecules by their characteristic absorption
spectra. Nucleic acids and proteins have specific absorbances at 260 nm and 280 nm,
respectively, and the measurement at these wavelengths and determination of the ratio
(260/280) will give the relative amounts of nucleic acid to protein in a sample.
Spectrophotometry can be used to measure the progress of a reaction and to characterize
solvents, chemical bonds and molecular groups.
Quantitative spectrophotometry is based upon the Beer-Lambert Law which
describes the relationship between a biomolecule and the amount of light it absorbs. The
Beer-Lambert law states that at a specific wavelength the fraction of light absorbed by a
given biomolecule in solution is proportional to the concentration of that biomolecule,
and to the path length of the light. This may be expressed in the following equation:
I
[Eq’n 1]
A  log 0  c
I
Where I0 is the incident light; I is the transmitted light;  is the extinction coefficient;  is
the path length, usually 1 cm; c is the sample concentration; and A is the absorbance of
the sample. The extinction coefficient (ε) is often called the molar extinction coefficient,
where c is in molar units.
A spectrophotometer does not give I0 or I, it gives the absorbance (A) or
transmittance. The relationship between absorbance and transmittance may be expressed
as:
A = log 1 = - log I
Eq’n. 2
T
Io
The absorbance is the most commonly used measurement.
According to Eq’n.1, when the Beer-Lambert law is obeyed, and absorbance is
plotted vs concentration; a straight line is obtained which passes through the origin. This
is a version of the equation of a straight line, y = mx + b, where b = 0, and = 1.
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Absorbance (A) has no units. It is related to the wavelength and written A280 , which
means the absorbance of the sample at a wavelength of 280 nm.
The extinction coefficients of different molecules may be very different, while
their wavelength maxima may be the same. The light absorbance of tryptophan is as
much as fourfold higher than that of tyrosine under identical conditions, however the
absorbance maxima for both amino acids occurs near a wavelength of 280 nm. Proteins
absorb strongly in the UV range of 280 nm, due to the presence of the amino acids
tyrosine and tryptophan. The proportions of these vary with the protein, and biological
samples are a mixture of many proteins.
The extinction coefficient (ε) is the absorbance of a unit solution and has units of
reciprocal concentration and path length. It is the relationship of the concentration of the
light absorbing compound and the path length of incident light to the absorbance of a
solution. This is usually a concentration of 1M and path length of 1 cm. ε600 = 4000 M1
cm-1 which means a 1M solution would have an absorbance at 600 nm of 4000 with a
path length of 1 cm.
Extinction coefficients are used for pure compounds only, and are used to
determine the purity and concentration of a compound, assuming it is soluble in aqueous
solution. ε values are specific for each compound, and knowing this from the literature,
and the concentration for the sample, one can determine the purity of the sample, i.e. how
close the measured ε is to the literature value. The higher ε is at a particular wavelength,
the greater the absorbance at that wavelength.
As there is no extinction coefficient for a mixture of proteins, one must use
another method to determine the protein concentration in a solution. Colorimetric assays
have been developed which use dyes to complex with the proteins, and these can be
easily measured. This only requires a visible light source. Many proteins have the same
response curve to these dyes, and the rapidity and stability of these assays make them
popular. If one needs to study a pure protein, that protein has to be extracted and purified
from the sample mixture.
A standard curve is a plot of absorbance vs. varying amounts of a substance. See
Standard Curves in the lab manual Introduction section for more information. A standard
curve must be linear over concentrations to be detected, and should not be extrapolated
for use past the highest or lowest standard values. The unknown concentration can be
determined from the subsequently generated graph. A reagent blank is necessary to
eliminate the absorbance from the sample solvent. The blank contains all the solvent
matrix, but no sample. The absorbance from the solvent is automatically subtracted from
the sample by the spectrophotometer when the blank is read and recorded.
When an unknown sample of a protein mixture is assayed, one usually makes
several dilutions (1/10, 1/100, etc) of the original sample to ensure that the concentration
will fall within the linear range of the standard curve. Do not forget to consider these
initial dilutions when calculating the concentration of the original sample.
One popular indirect method is the Bradford Dye Method which uses Coomassie
Blue dye in an acidic solution. The dye forms a complex with the proteins and is read at a
wavelength of 595 nm, which is in the visible range.
You will be constructing a standard curve for protein determination using this
method. The protein standard for the calibration curve is bovine serum albumin (BSA),
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very commonly used, and will be suitable for your calibrations. From your standard
curve, you will determine the unknown concentration of a protein that you will be given.
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Turn on the spectrophotometer as directed and allow to warm up while you are preparing.
This is usually 30 min. See Appendix C for instructions on using the Ultrospec 2100
pro Spectrophotometer
Lab Safety
PPE: Wear lab coat, goggles, and gloves
Chemical hazard: Low, you will use Tris, KH2PO4 and Bradford dye
Biological hazard: None
Physical Hazard: None
Solutions
o 20 mM, pH 5.6, KH2PO4
o 20mM Tris buffer pH 7.5
o Bradford assay
o MilliQ
Location
Cart
Cart
Cart
Back bench of lab
Exercise 7.1: Protein concentration and extract purity using
the Beer-Lambert law (direct assay)
**Keep your protein fractions on ice when thawed**
1. Set the wave length of the spectrophotometer at 417nm.
2. Take out your protein samples (from labs 5 & 6) from the ice bucket at the front of
the lab.
3. Keep all your fractions on ice during the lab.
4. Place a cuvette containing either buffer A or B and set the reference on your
spectrophotometer.
The type of buffer to use as blank depends on what your
proteins are dissolved in (see labs 5 & 6).
5. Make 1/10 dilution for all four protein fractions from labs 5 & 6. You will only need
a total volume of 1mL.
6. For each of the four fractions, transfer 1mL of your dilution into a disposable cuvette
and read the absorbencies on the spectrophotometer at 417 nm.
7. Using the Beer – Lambert equation, calculate the concentration of the myoglobin in
your fractions solution.
A= εc
where A= absorbance at the highest peak
ε = Extinction coefficient (7.57 ml. mg-1. cm-1)
 = path length (1cm)
c = concentration of protein
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Exercise 7.2: Determination of protein concentration by the
Bradford Protein Assay (indirect assay)
You will be constructing a standard curve of the protein Bovine Serum Albumin (BSA)
with known concentrations and using the commercial Bradford Dye reagent. The dye
reacts with all the proteins in the solution proportional to the concentration of the total
protein. From this curve, you will determine the concentration of the protein in your
samples.
1. Set the wavelength to 595 nm.
2. Obtain the following:
a. Six milliliter sample of the standard protein solution (0.1mg/ml BSA )
b. 20 mL Bradford dye solution from the stock by pouring into a small
beaker.
c. 72 eppendorf tubes.
3. For your Standard curve:
a. You are preparing triplicate samples of 8 different protein
concentrations, therefore you will need 24 eppendorf tubes. Label the
tubes 1.1, 1.2, 1.3, 2.1, 2.2, 2.3, etc. The first digit indicates the tube
number and the second digit represents the replicate number. Make
only one tube for the blank.
4. Using the most appropriate pipettor, pipette the required amount of standard into
each microfuge tube and dilute each to 800 μl with deionized water. See table
7.1.
Table 7.1
Tube No.
Reagent Vol (μl)
BSA μg
In
BSA standard
H2O
Bradford Dye
Triplicate
1
0.0
800
200
2
50
750
200
3
100
700
200
4
150
650
200
5
200
600
200
6
300
500
200
7
400
400
200
8
500
300
200
5. Calculate the amount of BSA in each tube as outlined in Table 7.1. Do this
before coming to the lab. Also show a detailed sample calculation in your lab
notebook.
6. For your four protein fractions (from Labs 5 & 6):
Make three dilutions of 1/5, 1/10 and 1/100 for each fraction. You will
need a total of 800uL for each dilution. You will need a total of 12 test
tubes (three dilutions for the four protein fractions)
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7.
Label an additional nine eppendorf tubes 1/5-1, 1/5-2, 1/5-3, 1/10-1, 1/10-2,
1/10-3, 1/100-1, 1/100-2 and 1/100-3 for each of the four protein fractions. In the
end you will have nine test tubes (3 dilutions) for each protein fraction.
8. Pipette 200 ul of your diluted protein sample (prepared in step 6) into the
corresponding eppendorf tubes from step 7.
9. Add to each tube (prepared in step 7) 600 μl of deionized water and 200 μl
Bradford dye reagent to each tube.
10. Incubate for 15 min at room temperature before reading at 595 nm. The reaction
is stable for 1 hour.
11. Zero or normalize the spectrophotometer with your blank, tube #1, to correct for
the absorbance due to the dye reagent solvent, by placing in the reference sample
holder. An absorbance of 0.0000 should be displayed.
12. Read the absorbance of each tube, starting from the lowest to the highest
concentrations.
13. Record all data, i.e. volumes and absorbencies
14. When you are finished, you should have all absorbencies in triplicate.
Lab clean up
1. Wash all glassware and test tubes, rinse, and dry and place back on cart or into your
locker
2. Test tubes used during the Bradford assay needs to washed with ethanol to remove all
the blue dye.
Exercise 7.3: Data Analysis for lab 7
This data analysis has to be performed before you come to lab 8. In addition bring
all your data to lab 8.
1. Calculate the concentration of myoglobin in your four fractions.
2. Calculate the average, standard deviation, and coefficient of variation of the three
absorbance readings for each standard.
3. Plot the average data on a graph of absorbance (y –axis) vs. μg protein (x-axis).
This is your standard curve. The graph should be linear over the
0-30 g range. The absorbencies of your samples should fall in
the linear range to calculate an accurate concentration.
Remember to include error bars.
4. Which dilutions (1/5, 1/10 or 1/100) can you use to determine the concentration of the
total protein in your four fractions. Why?
5. Using the standard curve, determine the concentration of total protein in the four
fractions from Lab 7.
6. Next week you will run your protein samples on a SDS-PAGE gel. To visualize your
protein on the gel you will need approximately 100 μg of total protein in a volume of
10μl (10 μg/μl). Before you come to class next week you will need to know how much
sample you need to load on your gels.
7. Determine the total protein and total Mb in the fractions. (amount not concentration)
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8. Determine the relative purity (RP) of myoglobin for all four proteins fractions. This is
defined as the total mg of Mb divided by the total mg of protein.
RP =
mg Mb
x100
mg total protein
9. How much protein and myoglobin was extracted into the 10 mL from Exercise 5.3.?
Lab 7: Lab Notebook Report
Your report will consist of the following:
1. Your report will consist of the duplicate pages from your lab notebook. Please read
the instructions on page 7. You will hand in the duplicate pages at the end of the
lab before you leave.
2. Answer all questions from exercise 7.3 in point form. This will be handed in next
week.
3. The standard (calibration curve) for the Bradford assay done using Excel. Remember
to show error bars, y-axis and x-axis labels etc. This will be handed in next week.
4. Table done in Excel showing standard deviation, average and coefficient of variation
for the absorbance readings. This will be handed in next week.
5. Table done in Excel showing the myoglobin and total protein concentration, total
amount of myoglobin and total protein and the relative purity of myoglobin in the
different fractions. This will be handed in next week.
As a reminder, it would be good to start with the formal lab report early (Labs 5, 6,
7, 8 and 9 combined). You can already start writing the Introduction, and some of
the Material and Methods and the Results section
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Lab 8: Characterization of Myoglobin by Sodium Dodecyl
Sulfate – PolyAcrylamide Gel Electrophoresis (SDS-PAGE)
Learning Outcomes
o To further investigate the protein myoglobin by the use of electrophoresis with SDSPAGE.
o To characterize the protein for both purity and molecular weight.
Introduction
One method which allows for both the visualization of the proteins within a
sample and a determination of their molecular weight is sodium dodecyl sulfate –
polyacrylamide gel electrophoresis (SDS-PAGE). Electrophoresis is the movement of a
charged particle within an electric field. The movement is dependent upon size, shape,
charge and chemical composition of the molecule. While electrophoresis can be utilized
to examine almost any biomolecule, it is especially useful in the separation of proteins
and nucleic acids. A typical electrophoresis setup utilizes a buffer saturated matrix on
which the sample to be analyzed is placed, and the molecules migrate through the matrix
due to an applied electrical field. The movement of these charge molecules is dependent
upon the electric field (E), the net charge on the molecule (q), and the frictional
coefficient (f). So the velocity at which the particle moves () is defined as:
Eq
Eq’n.1

f
The frictional coefficient depends upon the mass and shape of the molecule. The
electrophoretic mobility  is defined as

q
Eq’n. 2
 


E
F
Thus, the electrophoretic mobility depends solely on q/f, and is independent of the field
that is applied when the voltage is held constant, though it still relies upon the charge of
the molecule and its size. In most electrophoretic experiments the voltage is held
constant, though one can holdcurrent (voltage changes with resistance) or power ( the
product of voltage and current) constant if desired for the experiment.
Polyacrylamide gel electrophoresis (PAGE) utilizes a matrix formed by the
polymerization of acrylamide and has several advantages over other matrices. It has the
resolving power for small to moderately sized proteins, can accept large sample sizes, has
minimal interactions with the molecules, and is physically stable. Polyacrylamide gels
are generated by the free radical polymerization of acrylamide and its cross linker, N,N'methylene-bis-acrylamide (bisacrylamide), which is initiated by the
catalysts of ammonium persulfate
(APS) and TEMED (N,N,N'N'tetramethylethylenediamine) (Figure
9.1). More specifically, free radicals
generated by chemical decomposition
of ammonium persulphate and
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Figure 9.1. Polymerization
of acrylamide
and bis-acrylamide to
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TEMED are responsible for polymerization. APS generates SO4- free radicals when
dissolved in water. Therefore, it is always a good idea to make APS fresh since the free
radicals will decompose over time. The SO4- reacts with TEMED
(tetramethylethylenediamine) to produce TEMED free radicals that in turn induce
polymerization of acrylamide and bisacrylamide. Polymerization occurs in the absence
of oxygen since oxygen is a potent free radical scavenger. This is also the reason for
pouring acrylamide gels in between two glass plates to limited exposure to oxygen. The
degree of crosslinking is controlled by the relative percentages of acrylamide and bisacrylamide utilized, and the set resolving power and molecular sizes range of a particular
gel.
Electrophoresis depends upon the charge of a molecule, however proteins
normally do not have a uniform net charge associated with them. If the proteins are
subjected to treatment to give a uniform net charge, then their mobility will depend upon
their size, and will allow for the estimation of their molecular weight. Protein samples
are treated with the ionic detergent sodium dodecyl sulfate (SDS) which, in conjunction
with boiling and treatment with -mercaptoethanol, generates a denatured polypeptide
chain coated with negatively charged SDS and masking any native charges of the protein.
The overall negative charge will move the molecules toward the anode. Thus, a constant
charge/mass ratio is generated on all the proteins in the sample, and separation will be
dependent upon the sieving of the gel matrix based on a molecular sieving (gel filtration)
effect.
Discontinous gel electrophoresis is a further refinement of the electrophoretic
process, in which there are two gel layers of differing cross-linking content, ionic
strength and pH, and the upper layer (stacking gel) has a lower percentage cross-linking.
These differences allow the formation of a concentrated sample band which allows for
greater resolution of the sample in the main (resolving) gel.
Once the proteins have been separated in the gel by electrophoresis, the proteins
are subject to diffusion away from their final position when the electric force is removed,
so they must be fixed in place. In addition, in order to be analyzed the proteins must be
made visible. Fixing of the protein position and visualization are two steps normally
performed in conjunction with one another. Two simple methods for visualizing the gel
are dye staining, and deposition of silver (Ag) at the protein band. Coomassie Brilliant
Blue is a dye which non-specifically interacts with proteins, and is the most commonly
utilized staining procedure for bands.
Coomassie staining can detect 0.5 to 1 g of
protein per band. Once the whole gel is saturated with dye, the gel is washed, removing
the dye from all but the protein bands. Silver staining is the result of the reduction of
ionic silver (Ag+) to metallic silver, where the difference between the reduction potential
of the protein and the acrylamide results in a larger deposition of metallic silver at the
protein bands. Silver staining is more sensitive than Coomassie staining as it can detect
nanogram quantities of protein per band. You will be staining your samples with
Coomassie Blue for visualization in this experiment.
To utilize electrophoresis to determine the molecular weight of a sample, its
mobility has to be compared to the mobility of a set of proteins of known molecular
weight, utilizing a standard curve. As the movement of these standards depends upon the
exact conditions of the experiment, they must be included in every electrophoresis gel
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run. The mobility of a protein is dependant upon its size and is related to the distance the
protein moves in a gel, also known as the retention factor (Rf).
migration of band
R f  retention factor 
Eq’n. 3
migration of dye front
The purity of a sample can also be determined through electrophoresis by examining the
number of bands that appear in the developed electropherogram (the developed gel). If
more than one band is present, then the sample is said to be heterogeneous, while if only
 one band is present, the sample is homogeneous.
In today's experiment, we will be performing sodium dodecyl sulfate –
polyacrylamide gel electrophoresis (SDS-PAGE) on your extracted myglobin and
fractions taken during the purification, see Lab 6. The electrophoresis experiment serves
several purposes. First, it allows you to monitor the effectiveness of the purification steps
you carried out last week. Second, it allows you to determine which fractions from your
columns in Lab 6 contain myglobin and third, it allows you to estimate the molecular
mass of your purified myglobin.
Lab Timing
o Start immediately with Exercise 8.2 and 8.3
o Pre-lab Talk while performing the electrophoresis
o Exercise 8.4
Lab Safety
o PPE: Wear lab coat, goggles, and gloves
o Chemical hazard: Acrylamide is a potent neurotoxin. You will also heat acetic acid
and methanol. The vapours from acetic acid and methanol can be quite
overpowering. Therefore work with these chemicals in a fume hood.
o Biological hazard: None
o Physical Hazard: High voltage electrical current from electrophoresis apparatus.
Solutions required
o 10x Tris/glycine running buffer
Label
--
o 2x Gel loading dye
o -Mercaptoethanol
o Staining solution
2X LD
B -Me
--
o Destaining solution
--
Location
Electrophoresis
bench
cart
fumehood
Electrophoresis
bench
Electrophoresis
bench
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Exercise 8.1: Preparing SDS-PAGE using the BioRad MiniProtean System
Exercise 8.1 was prepared for you, but you are still required to be familiar with the
theory.
The procedure you will use is essentially that originally published by Laemmli.
You will be using the Bio-Rad Mini Protean11. There will be 2 gels made per apparatus,
and one gel per group. You will be running your experimental samples as well as low
molecular weight standards and a myoglobin standard.
1. Make sure that the glass plates are clean and dry before use. Always wear gloves
when making up your gels, solutions and loading your samples. Assemble the
apparatus before making the solutions below.
2. Assemble the casting stand and plates as instructed. Make your resolving (lower)
gel as described. Unpolymerized acrylamide is a potent neurotoxin. Wear
gloves and use pipette aids when handling gel solutions.
3. To make you Resolving gel of 15% monomer, mix the following in a small
beaker (Total volume will be 6 mL). Mix well, but do not shake to prevent
bubbles forming due to the SDS:
30% acrylamide solution
3.12 ml
Lower gel buffer
1.5 ml
Deionized water
1.38 ml
4. To 5 ml of the above solution, add the following and mix briefly:
20% APS
10 μl
TEMED
3 μl
5. Pipette approximately 4 ml of this solution between the glass plates in the
apparatus up to a previously marked level to make room for the stacking gel.
6. Add approximately 200 μl of methanol or saturated butanol to “flatten” the top of
the gel while it is setting. It will take about 30 min for the gel to fully polymerize.
Since aqueous solution will mix with the gel organic solvents such
as methanol or butanol are used to flatten the gel. Nevertheless
a small amount of water is added to the organic solvent since
pure organic solvent will dehydrate the gel resulting in shrinkage.
7. When gel has polymerized decant butanol, and wash the gel carefully with
deionized water using a pipet and dry the plates as much as possible using the
corner of a KimWipe.
8. Make your stacking (upper) gel as described below and pour over the lower gel.
Insert the required comb and allow it to set.
i. Stacking gel – 2.007 ml of 4% monomer
a. Stack mix
2 ml
b. 20% APS
5 μl
c. TEMED
2 μl
ii. Mix briefly and immediately add this solution to the top of the
resolving gel.
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iii. Insert the desired comb into the stacking gel slowly, avoiding
bubble formation.
iv. It will take about 30 min for the stacking gel to set.
9. Remove the gel from the casting stand as instructed. Gently remove the comb
from between the plates.
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Exercise 8.2: Protein Quantification and Sample Preparation
1. To visualize your protein on the gel you will need approximately 100 μg of total
protein in a volume of 10μL (10 μg/μL). Dilute your protein samples to the desired
concentration and make sure you have at least 30 μL. If the concentration of your
samples are less than 10 μg/μL dilute the crude extract and filtrate fractions 1/10
and the protein fractions (those in Buffer A and B) from the column chromatography
are used without dilution.
Perform the following in a Fumehood
2. To 200 μL of 2X loading dye add 20 uL of β-mercaptoethanol.
Loading dye contains a number of components. SDS and DTT
denature secondary and tertiary protein structures to yield
proteins in the form of extended negatively charged rods. This
is promoted by boiling of the sample with the loading dye. BMercaptoethanol is added to reduce di-sulfide bonds in the
tertiary structure of the protein. EDTA is included in the dye
to chelate metal ions to prevent degradation of your sample by
metal dependent proteolytic enzyme. Lastly, glycerol is added to
pull sample down into the well and bromophenol blue is used as
tracking dye.
3. For each protein fraction, pipet 20 μl of sample ( prepared in step 1) and 20 μl of
loading dye into a 1.5 ml microfuge tube.
4. Pipet 20 μl of the myoglobin (Mb) standard and 20 μl of loading dye into a 1.5 ml
microcentrifuge tube.
5. Your Low Molecular Weight marker (Protein ladder, Appendix D) already contains
loading dye just add 1 μl β-mercaptoethanol.
6. Pulse all the tubes for 10 seconds in a centrifuge
By doing a quick centrifugation step the sample and dye is
properly mixed and forced to the bottom of the tube
7. Boil your samples, Mb standard and protein ladder for 5 minutes in the boiling water
bath located in the fume hoods
8. Cool and pulse spin the tubes (10 seconds in the centrifuge). You are now ready to
load the samples on the gel
Exercise 8.3: Running SDS-Page Gel
For this exercise two groups will work together.
1. Make 500 mL 1 x Tris/Glycine from the 10 x Tris/Glycine and pour into a 600 mL
beaker until use. Do not mix this too vigorously as foam is readily formed from the
SDS content of the buffer.
2. Each group will have one gel with two gels per electrophoresis system.
3. Assemble the Mini-Protean II following the instructions. Fill the buffer chamber with
1X Running Buffer as demonstrated.
4.
Load samples in gel according to the following pattern:
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Well #
1
Sample
I.D.
LMW
Std.
2
Mb
Std.
Biochemistry Laboratory
3
4
5
Crude
Extract 1
Filtrate 2
Buffer A
Sample
6
Buffer B
Sample
5. This procedure will be demonstrated. Using a gel loading tip, carefully pipette up
to 20 μL of standards into the appropriate wells. Next load the samples into the
respective wells using a new gel loading tip for each sample. Load the protein ladder
last.
This procedure must be done fairly rapidly to avoid lateral diffusion of
the proteins into the gel matrix, but don't rush and mix up or overflow
the wells.
6. Securely place the lid on the electrophoresis apparatus and connect the leads to the
power supply.
7. Set the power supply to 240 V, constant voltage.
8. Electrophorese the gel for 60 min or until the bromophenol blue dye is within 0.5 to 1
cm of the bottom of the gel.
This will be demonstrated in the lab. Be careful not to touch the
surface of the gel as proteins from your hands will transfer and
contaminate it.
9. Cut a notch in the gel, along one edge, at the point marking the distance migrated by
the tracking dye.
Excess, polymerized gel can be disposed off in the hazardous solid waste
container ( yellow bucket).
10. Proceed immediately to Exercise 8.4 to stain the gel.
11. The electrophoresis unit must be washed and left to dry on the peg rack. Carefully
wash the glass plates with soap and water and leave them to dry on the finger rack
provided.
Exercise 8.4: Staining the Gel with Coomassie Blue
1. After electrophoresis, the gel is removed and placed in staining solution of 0.04%
w/v Coomassie Blue G-250 and 10% v/v acetic acid and 40% v/v methanol.
2. Microwave for 45 seconds or until solution starts to boil. Be careful when
removing the container from the microwave. The solution contains acetic acid
3. Pour stain back into bottle for later use.
4. Add destaining solution (10%acetic acid 40% methanol) and heat in microwave
for 45 seconds or until boiling. Change destain solution and incubate at room
temperature until the proteins are detected as dark blue bands.
5. Identify the bands that you see in your ladder and identify the protein samples
that contain myoglobin.
6. Using the proteins of your Low Molecular Weight marker plot a curve of the Log
MW vs. Retention factor.
7. Determine MW of myoglobin.
Lab clean up
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1. Wash all glassware and dry and place back on cart or back into your locker
2. Refill your tip boxes
3. Take gel apparatus apart, wash, rinse with distilled water and dry
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Biochemistry Laboratory
Lab 9: Atomic Absorption Spectrometry of Iron content in
ground beef
Learning Outcomes
o To develop an understanding of Atomic Absorption Spectrophotometery
o To describe the importance of inorganic chemistry in biochemistry
o To relate the Iron content to myoglobin content in ground beef
o To construct a calibration curve
Introduction
Atomic absorption spectroscopy (AAS) is a qualitative and quantitative analytical
technique that measure ultraviolet and visible light radiation from elements in their
gaseous state. AAS has many applications and proved to be a reliable analytical
technique in determining the amount and type of elements present in blood samples, soil,
milk, and many food products. There are two main types of AAS: flame atomization and
electrothermal atomization. In this lab you will be using a flame atomization
spectrometer. A typical flame atomization AAS uses a flame that vaporizes a solution
(usually aqueous) containing the element you are interested in analyzing (Fig. 9.1).
Fig. 9.1. A typical single beam Atomic Absorption Spectrometer.
The element in its gaseous phase will then absorb radiation from the cathode tube (light
source) (Fig. 9.1). The light absorbed by the element is directionally proportional to the
concentration of the element. This is be described by the Beer’s Law (see also lab 7 for
more details on Beer’s law):
A = log 1 = - log I
Eq’n. 1
T
Io
To measure the intensity of the absorption by the element, excess light travels through a
monochromator that is set to allow only light of a certain wavelength to pass. Every
element has a characteristic absorbance, meaning that the element will absorb light at a
specific wavelength. The monochromator is set by the operator depending on the
element being analyzed. The light that passes through the monochromator is then
measured by a detector and converted into signal readout.
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Biochemistry Laboratory
**After completing Lab 9 you will perform Exercises 10.1 and
10.2 as well.**
Lab Safety
PPE: None.
Chemical hazard: None.
Biological hazard: None.
Physical Hazard: None.
Exercise 9.1: Atomic Absorption Spectrophotometry analysis
of Iron extract from ground beef.
1. Your samples will be placed in the autosampler of the AA. Currently your
samples are labeled as Sample01, Sample02 etc. To change labels to your sample
names click on the “Labels” tab and enter the new names
2. When you are ready to run your samples make sure you are in the “Analysis” tab.
3. Press “Select” and then highlight the samples you would like to run. Press
“select” again. When you are ready to acquire the data, click on the “Start”
button.
This will cause the AAS to prepare and zero the instrument, and run your samples
in order. The absorbance values will be recorded in each cell. You will also see a
live plot of each absorbance being acquired on the right hand side of the screen.
Clicking on overlay at the bottom of the plot will allow each to be compared. It
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Biochemistry Laboratory
takes longer to acquire low concentration samples or ones with a lot of noise.
Record the absorbance values in your lab notebook once the readings are
complete.
4. Repeat steps 1 to 3 twice so that you have you data in duplicate
5. The concentration of iron in your standards are 5 mg/L, 10 mg/L, 25 mg/L,
50 mg/L, and 100 mg/L.
**After completing Lab 9 you will perform Exercises 10.1 and
10.2 as well.**
Exercise 9.2: Data analysis
1. Calculate the average, standard deviation, and coefficient of variation of the
three absorbance readings for each standard and the sample.
2. Plot the average data on a graph of absorbance (y –axis) vs. iron concentration
(x-axis). Remember to include error bars.
3. Using the standard curve, determine the concentration of iron in your original
sample from lab 5.
4.
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Biochemistry Laboratory
Labs 5, 6, 7, 8 and 9: Formal Lab Report
Your report will be evaluated for overall quality of the writing, grammar, spelling, the
ability to construct tables and figures (graphs) correctly and the ability to analyze and
interpret data appropriately. Please read the guidelines on writing formal lab reports in
the beginning of this manual. Include all the required data. Be sure to answer all
questions from labs 5, 6, 7, 8 and 9. The answers to these questions, depending on the
question, should be incorporated as part of your result or discussion section. You are
allowed a maximum of 15 pages. Title page and appendix are excluded.
For your formal lab report include the following sections:
Report (90) + lab demeanor (10) =100
o Introduction (10)
o Results (45)
o Construct a double plot of absorbance at 417nm and 280 nm against fraction
number
o Report total protein concentrations
o Concentration of myoglobin in each of the samples
o Relative purity of myoglobin in each of the samples
o Show Bradford assay standard curve
o Photo of SDS-PAGE gel
o Standard curve of log (MW ) and Rf
o Molecular Weight (MW) of myoglobin
o Atomic Absorption standard curve
o Moles of Iron and myoglobin in the 10mL of exercise 5.3
o Discussion (25)
Questions for Discussion
o Why do we measure myoglobin at 417nm?
o At what pH will the myoglobin bind to the column? Why?
o How much of the myoglobin you loaded onto the column did you recover?
Calculate the percentage recovered. If not all of the myoglobin was
recovered what could be reasons for the loss of protein and how would you
improve your recovery?
o How do the MW of myoglobin from your SDS-PAGE compare to published
literature values?
o There should be a one-to-one stoichiometric relationship between iron and
myoglobin. Try to correlate the amount of myoglobin and iron you have, do
you observe this stoichiometry? If not, why is there not stoichiometry?
o References (3)
o Appendix (7)
o Any sample calculations from labs 5 to 9. Make sure you write out in full
detail the whole calculation.
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Commons, 171 Second Street, Suite 300, San Francisco, California 94105, USA.