PURIFICATION OF -GALACTOSIDASE FROM E coli.
for Biomolecular Techniques: Proteins
I.
Introduction
A.
Background
B.
Overview of Project
C.
Written Report
D.
Laboratory Notebook
II.
Protocols
A.
Buffers
B.
Extraction
C.
Ammonium Sulfate Precipitation
D.
Initial Assays
E.
Dialysis: to Desalt or Change buffer
F.
Ion Exchange Chromatography
G.
Analysis of Column
H.
Final Assays
III.
Technical Notes
A.
Cold
B.
Reducing Agents
C.
Protein Assays – issues of particular note
D.
Column Selection and other considerations for the column
IV.
Calculations
*************************
I. INTRODUCTION
This project was originally developed from lab # 57 in Experiments in Molecular
Genetics, Jeffrey H. Miller, Cold Spring Harbor Laboratory, 1972. It was adapted for use in
the Protein Bioseparations course in the Biotechnology Laboratory Technician Program at
Madison Area Technical College (Wisconsin) by Lisa Seidman, Ph.D and has been further
modified to reflect the equipment and experiences of the Biotechnology program at Portland
Community College.
A.
BACKGROUND: -GALACTOSIDASE
The cells of microorganisms usually have a wide variety of enzymes. Often certain of these
enzymes have the property of being inducible; that is, their concentration will increase if their
inducing agent, often their substrate, is present. The enzyme -galactosidase, found in E. coli,
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is such an enzyme. This enzyme hydrolyzes a wide variety of sugars (those disaccharides or
oligosaccharides which have a linked galactose residue in their structure). When normal
cells are grown in the absence of lactose or other -galactosides (for example, when glucose is
a major carbon source) -galactosidase is present in very low levels. This is because a
repressor prevents the transcription of mRNA for the enzyme. When a -galactoside such as
lactose is present in the medium, the repression is turned off and production of the enzyme
increases as much as 1000 fold.
In some strains, -galactosidase is produced at higher than basal levels even in the absence of
the inducer. Such strains are called “constitutive” (this usually results from a mutation
which prevents full repression). In our experiment, we will be using a strain that is a
constitutive -gal producer.
The -gal enzyme is not only expressed at high levels in this strain, but it is in general a pretty
sturdy enzyme. Its activity is easy to measure, because there is a substrate, which, when
cleaved, turns colored, so that the progress of the reaction (generation of product) can be
quantitated. It can be purified quite well by just a couple of steps and is a good choice for
learning some key protein techniques.
B.
OVERVIEW OF THE PROJECT
After preparing a crude cell extract, we will carry out a traditional sort of purification
scheme, starting with a bulk precipitation procedure, followed by ion exchange
chromatography. With a combination of enzyme and protein assays), we will evaluate the
success at each step (how much enzyme did we recover, and how much more pure is it?). We
will also evaluate the protein composition at each step by separating the proteins on a gel
(SDS-PAGE) and staining.
C.
WRITTEN REPORT
Formal communication of scientific work is generally done in the form of papers that are
submitted to a peer-reviewed journal, for publication intended mostly for others in the field.
Because there is this expectation that the work will be read mostly by other scientists, it can be
difficult to get used to the format and style of a standard scientific publication. The good
news is that there IS a standard format and style, and many conventions that make it easier for
us to communicate with one another.
To help you learn how to work with communication in this format, you will write up each part
of this project as though it were being presented for publication, and by the end of the term,
you should have a paper which summarizes the work you did, the data you obtained and
highlight the meaning and importance of the results. There is a separate handout for this.
NOTE: Although you will be working in a group, the papers will be written independently.
The data will be the same for each member of the group, but I expect that the presentation of
the data – the write up, the tables and graphs, to be the work of each individual.
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D. LABORATORY NOTEBOOKS and working in groups
The expectation of a written document tat the end does not get you off the hook for
maintaining a detailed laboratory notebook! In fact, it is just the opposite – you cannot hope
to prepare such a paper of you lack the important records of what you did and what the results
were. Remember to be especially vigilant about details such as the volumes you have at each
step, the location where things (such as your various fractions!) are stored, the times for
various incubations, centrifuge spins etc. Never throw away any supernatant, pellet,
fraction or sample until the entire project is complete.
Group work presents particular challenges for maintaining a smooth record of events. In a
project like this one, where you will be responsible for reporting everything that happened,
and every bit of the results, you will need to make sure you fill in your notebooks with results
from your group members. You can photocopy and tape in, or you can write out information
obtained from their notebooks – in either case, be sure to attribute it to the person who
actually did the work.
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II.
PROTOCOLS FOR PURIFICATION OF  GALACTOSIDASE
Each team will start with some volume of bacterial culture (our constitutive -gal producer
ATCC 15224) – or a pellet derived from such a culture
A.
SOLUTIONS
Breaking buffer
0.2 NTM
0.4 NTM
0.2 M NaCl
0.2 M Tris, pH 7.5
10 mM Mg acetate
5 mM Dithiothreitol (DTT)
5 % (v/v) glycerol
0.2 M Na Cl
10 mM Tris, pH 7.5
10 mM Mg Acetate
5 mM DTT
0.4 M Na Cl
10 mM Tris, pH 7.5
10 mM Mg Acetate
5 mM DTT
note for all: add DTT
just before using
B.
EXTRACTION
1. Harvest bacteria by centrifugation in pre-weighed 250 mL bottles at 6000 rpm (GSA
rotor), for 10 minutes.
2. Resuspend the cells in Breaking Buffer. Use a volume equivalent to 5 mL per gram of
pellet weight (expecting 5-7 grams of cell pellet per liter of culture) in mL of Breaking
Buffer.
3. Set up ring stand, sonicator and ice bath to hold the beaker with your cell suspension.
4. Sonicate, on ice, at 50% power setting for 10 minutes (2 sec pulse).
5. Centrifuge at 10,000 x g for 30 minutes
6. Remove the supernatant (= starting lysate, or extract) , record the volume
7. Reserve 1-2 mL of the extract for later analysis
C.
AMMONIUM SULFATE PRECIPITATION
Previous data suggested that 45% saturated ammonium sulfate is appropriate to precipitate
most of the -gal, but less than half of the total protein.
1. Pour the remaining supernatant into a beaker with a stir bar, allowing for a 50% volume
increase. Set stirring in the cold.
2. Measure the pH. (Remember to measure it cold, because Tris is temperature-sensitive)
3. Determine and measure out the necessary amount of ammonium sulfate to bring this
volume to 45% saturation (at 4 C).
4. Add ammonium sulfate slowly, over a period of 30-45 minutes, with gentle stirring.
4
5. Measure the pH. (ammonium sulfate tends to acidify the solution) The pH should be
between 6 and 8. If it is not, adjust it with NaOH or HCl.
6. Continue stirring in the cold for 1-2 hrs (overnight is OK, but 1-2 hrs seems to work
better)
7. Centrifuge the suspension at 10,000 x g for 10 minutes.
8. Decant (and save) the supernatant (store at 4 C, carefully labeled, including the volume!)
9. Resuspend the pellet in some volume of 0.2 M NTM
We are expecting our activity to have precipitated – putting it in a smaller volume
allows us to concentrate it, so that we don’t have so much volume to load on the
column, but in fact, the volume is not really critical here. And we want to have
enough to handle – we have to dialyze it first, and very small volumes are hard to
dialyze. Probably between 5 and 10 mL is good
10. Remove and reserve a small volume of the sup and resuspended pellet for later assay
(reserve no more than 10% of the total. This is just lost, as far as the purification is
concerned. Don’t lose more than you need to.)
D. INITIAL ASSAYS
At this point we will do enzyme and protein assays, to make sure we are proceeding with the
correct fraction. However, before we can load the column, we need to dialyze, so that the
sample is in the correct salt concentration to bind to the ion-exchange column. This is not a
purification step, but we might lose protein or enzyme activity or both, so it would be good to
assay the fractions both before and after dialysis
See separate handouts for:
BioRad Protein Assay
-galactosidase Assay
also, see the section on CALCUATIONS, below
E.
DESALTING AND CHANGING BUFFER BY DIALYSIS
1. Handle dialysis tubing with gloves. Do not let it dry out.
2. Cut a piece of dialysis tubing sufficient to hold your resuspended pellet.
3. It is good to soak the tubing in water, stirring in the cold, for at two hours. This will
remove impurities, such as antifungal and antibacterial agents, with which the tubing may
have been treated. Some labs autoclave the tubing in EDTA – this is mostly useful for
working with DNA (kills DNases!).
4. Remove the water carefully from the tubing. Tie a knot or two, or clip the bottom
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5. Carefully pipette your sample into the dialysis tubing, and tie it off (leave a bit of slack
for increase in volume due to osmosis. I like to make sure there is a bit of air in there so
that the tube floats and doesn’t get bashed about with the stir bar)

Keep track of your volumes so changes due to osmosis can be corrected for !!!

Make sure that you keep the sup and pellet clearly marked! .

Dialysis could result in other changes, such as formation of an insoluble
precipitate, or loss of proteins that are smaller than the MWCO of the dialysis
tubing, so you will want to assay both pre- and post-dialysis samples to compare.
6. Dialyze against 1000 mL of 0.2 M NTM buffer, 4 hrs or longer)
7. Change the dialysis buffer (another 1000 mL of 0.2 M NTM at least twice before the next
class.
8. Remove the dialyzed samples from the tubing into a centrifuge tube.
9. If the samples are not completely clear, centrifuge 10 minutes at 10,000 x g to remove any
insoluble material. (Don’t try to recover this pellet for assay, but note whether there was a
pellet, because it may help to account losses in activity or protein)
10. Record the volumes.
11. Check the pH of the dialysate. This is actually important. It must be the same as the pH of
the 0.2 NTM buffer (this is so that the ion exchange conditions will be correct). If it is
not, adjust it.
F.
ION EXCHANGE CHROMATOGRAPHY FOR -GALACTOSIDASE
1.
Prepare a DEAE Sepharose Fast Flow column. (see notes below)
2.
Equilibrate the column by washing with 0.2 M NTM until the pH and the conductivity
of the buffer and the effluent are very close. Note: for packing and equilibration, you
can use a flow rate that is just under the maximum for the resin.
3.
Change to a slow flow rate (1 mL/min or less; you want to give the sample plenty of
time to bind). Load the sample on the column, collecting the effluent in several large
fractions. Monitor the A280 so that you
a)
can locate the fractions containing most of the unbound protein, and
b)
Know when to begin the salt elution (when A280 has come down to zero)
You will want to assay the unbound fraction for protein and for enzyme
activity. You hope that lots of protein comes out here, and very little enzyme
activity (you want that to elute from the column, with less protein). Because
the salt concentration is relatively high (0.2 M NaCl) you expect many proteins
to not bind. We keep fractions so that we don’t dilute the unbound protein
unnecessarily. If we want to see what it looks like on a gel, we will need a high
enough concentration to load 10 ug in about 20 uL.
6
4.
You can assume that when the A280 of the column eluate has come down to baseline
(near 0.000), that the unbound proteins have been collected. Now you will elute the
column with a 0.2 M to 0.4 M NaCl gradient (in NTM) using between 4 and 6 column
volumes for the elution, collecting between 60 and 100 fractions. Elution can be done
at a much faster flow rate.
So if you have a column that has 20 mL of bed volume, the gradient might be
100 mL, which means 50 mL of each of the two solutions (the low and the high)
will be mixed using a gradient maker.
5.
“Step” off proteins that have not yet eluted with a higher salt concentration: 1 M or so
(in NTM). One column volume should be adequate, and if you wish to reduce the
number of tubes, you could collect it in larger fractions (maybe 4 or 5 for the whole
additional volume.
That’s all there is to it! But here there are a number of issues for you to consider – be sure to
check the section on Technical Notes.
G.
ANALYSIS OF THE COLUMN and POOLING THE KEY FRACTIONS.
1. Assay the fractions of the column for both protein and for enzyme. This is tedious but
important. (Note: sometimes people do every other fraction, or every third fraction, when
preparing a column profile. This is and acceptable practice, especially if you are
concerned about recovering the maximum amount of material, but you can miss things.
Because this is a learning experience, I would like you to assay each sample. It is not all
that hard!
Protein can be done by A280 (read samples directly in quartz cuvettes – which can even
be done while the column is running), or by the BioRad Protein Assay (single samples
are OK). In either case, it is important to go back over sections with very high
absorbances, and assay a smaller amount (for A280, you will need to dilute, since it
takes a minimum volume of 0.7 mL to read) to make sure you are in the linear range
(or in the case of the BioRad Protein assay, the “linear” range) of the assay [see below]
The enzyme assay can be done as single samples too. I would recommend going
through the whole column under a standard set of conditions (say 50 uL and 2
minutes). When you find the peak of activity, if the absorbances are too high (A420
greater that 1.0), re-do these sections with smaller volumes or dilutions of the fraction.
Try not to consume too much of your fractions in analysis – these procedures are
necessary, but do consume product – be judicious in their use.
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2. Calculate to plot the protein and enzyme concentration.
Excel can be very helpful here because it allows you to apply one calculation to a large
string of numbers, and bring in different volumes, dilution factors, times etc to each
one.
For the protein assay, you can use a calculated y = mx + b as long as all of the samples
are in the “linear” range of the assay. We can go over how to do this.
3. Plot the protein and enzyme concentrations vs. column fraction – superimposed on the
same graph. (If someone is better at Excel than I am, they can show us how to do this, but
if not, I would rather you just plot them by hand, as people did for years and years before
Excel! It is not as time consuming as it would seem!) Use different symbols or a different
line pattern for the two curves.
Note: you might think it would be just as good to plot the absorbances for each assay
on the curve. Why is this usually not OK? Under what conditions would it be OK?
(answer: if, and only if, all assays used the same volume/dilution of sample, and the
same time of incubation – which usually is not the case, if the assays were done
correctly)
4. Pool fractions. By examining the column profile you should be able to see groups of
fractions that seem to all contain the same protein, and can be combined (or “pooled”)
pooled together. The idea is to get the maximum amount of enzyme with the least
contamination from anything else, which is why you need to see BOTH the enzyme and
the protein profiles together. You also need to make decisions about how to balance the
desire for yield, and purity, and concentration (if you want to see the protein on a gel).
Choosing which fractions to pool is somewhat of an art, and depends very much on what
your data and your needs are. You should consult with the instructor about this before you
pool.
H.
FINAL ASSAYS
In addition to assaying the pools, you will need to repeat the enzyme assay on the stored
lysate, and on the material that went on the column (reserved dialyzed pellet, most likely).
The reason for repeating the assay it that the enzyme will likely have lost activity in the
intervening weeks. You want to know that, and also be able to compare, for example, how
many units went on to the column, in “today’s units” (that is, assuming that whatever loss in
activity the enzyme suffered over time is equivalent in the purified prep vs. the reserved lysate
– NOT a valid assumption, but the best we can do under the circumstances), vs. how many
came off. If you only had the prior assay, and you get off fewer than you put on, you would
not know whether you were looking at loss of enzyme molecules or loss of activity. Protein
assays are not as critical to repeat (unless wholesale degradation as occurred, they should be
the same.) (See calculation section)
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III.
IMPORTANT TECHNICAL NOTES
A.
COLD
Throughout this project, do your best keep the enzyme cold, either on ice or in the refrigerator.
The enzyme loses activity when it kept at warm temperatures for extended periods of time.
(Proteases present in the cell extract may cut at it, and even a pure protein can suffer activity
loss as a result of oxidation, broken H-bonds and other molecular events). Use ice baths,
refrigerated buffers, refrigerated centrifuges, etc. as much as possible throughout.
Experience suggests that freezing the enzyme, may prolong the activity, but we don’t have a
great deal of data to support this. It may depend on which buffer the enzyme is in, and the
time of storage. Repeated freezing and thawing usually does damage proteins (the ice crystals
can break bonds), so enzyme frozen for long term storage should always be aliquotted (so that
it is not repeatedly frozen and thawed. You are welcome to investigate appropriate storage of
the enzyme experimentally, if you wish.
B.
REDUCING AGENTS
- mercaptoethanol (BME) or dithiothreitol (DTT) are used throughout as reducing agents you
will note that the concentration is 5 mM DTT. This is not here to break disulfide bonds, but to
preserve the integrity of sulfhydryl and other oxidizable groups. In the organism, proteins are
in a reducing environment and oxidation causes problems (including the dreaded free
radicals). In the lab, in dilute solution, proteins are susceptible to oxidation, and this can
affect their structure and function. The concentration is a little higher than the concentration
typically used for protection of oxidizable groups, and suggests that there may be a sulfhydryl
group important for enzyme integrity.
Reducing agents do themselves become oxidized in dilute solution. This is, after all, how
they work -- by essentially out-competing the oxidizable groups on the protein, and thus
protecting them. At high concentrations (BME stock is liquid13.4 M, and DTT is stored as
a 1 M they remain reduced because these solutions are relatively acidic, and there is a lot of
H+ around! For this reason, it is important to add DTT or BME to your buffers just before
they are used. You can prepare the buffer without reducing agent ahead of time, with an
annotation that DTT has not yet been added.
DTT is a better reducing agent than BME. For that reason, BME is used at a higher
concentration than DTT. The protocol we are using calls for DTT at 5 mM. In “Z Buffer”
(universally used in b-galactosidase assay) BME is used at a concentration of 38.6 mM,
nearly 8 X higher than the 5 mM DTT called for in the buffers of the purification. I am not
sure why the concentration is so much higher, but go ahead & use the BME as indicated.
Both of these reagents smell strongly, so use hoods and closed containers whenever possible.
9
C. PROTEIN DETERMINATIONS and special issues for this project
For the most of our analytical work, we will do the BioRad Protein Assay, based on the
method of Bradford, and using BSA standard. There are other assays, notably the Lowry and
the BCA assays, that offer some advantages, but also have their own drawbacks. (See BioRad
Protein Microassay Protocol). It’s also true that you could use other protein standards.
Important note:
It is NOT A GOOD PRACTICE for one person to do the standard curve and another to do the
samples. This is just as bad a plan as doing the standard curve one day and the samples
another. Too many variables can affect the results. Find a way to share the workload without
invalidating the data. Also, please try to work efficiently. The tubes and cuvettes are
expensive, as is the reagent. Don’t do replicates until you know that you are in the right
range.
Protein Determination from Column Fractions
For most of the sections of this project there are just a few (under 20) sample, but for the
DEAE column, you will need to assay many samples (ideally, every fraction), in order to see
how protein eluted from the column.
There are two ways to do this:
If you want to, you can do the BioRad Protein assay reaction in 96 well plates. I would premix the Concentrated reagent, ¼ (1 mL CR + 3 mL water), put 160 uL of that into each well,
and add 40 uL of each fraction, triturating to mix. Alternatively, you can or do the reaction in
tubes and then transfer to 96 well plates to read. You should set up a standard curve, in
triplicate. (Samples from the column do not need to be done in triplicate)
Or, you can use a different method altogether read the absorbance at 280 nm. At this
wavelength, the amino acids that have rings (phe and try) absorb, and you can get a rough idea
of protein concentration. This is not good for crude extracts (too much RNA, which absorbs
maximally at 260, and significantly at 280), and it is not good to use this for following protein
in an assay, because proteins vary so widely in their amino acid composition, that it is only
good for a rough estimate.
A280 must be read in [very, very expensive] quartz cuvettes, so the samples are read one at a
time.
Interfering Substances in Protein Assays
Some buffer components can interfere with almost any assay you use, and lead to erroneous
results.
In the Bradford protein assay, for example:

Some substances can prevent (or attenuate) binding of CBB with the protein,
diminishing the signal and thus causing an underestimate of protein

Some substances can cause the absorbance shift to occur independent of protein,
and resulting an overestimation of protein in the sample
10
It is often possible to look up which substances have been found to interfere, and at what
concentrations, but you can also use an empirical approach. Before you kill yourself doing
this, find out if it will matter. In most cases, small amounts of sample contain insignificant
amounts of the potential interfering substance. If you find that you can get away with less than
50 uL in an assay, you may not need to worry about it. If you cannot tell by looking it up, try
this:
Make two tubes each of blank and 5 ug standard. To one pair, add the same volume of
interfering substance that you want to try (for example, if you are concerned that the salt that
is in a 50 uL of a sample in 0.4 M NTM will be a problem, add 50 uL of 0.4 NTM to one
blank, and also to one 5 ug tube. Then add the CR, vortex and read. Are they different? (Do
replicates if you want to be sure)
If the buffer does seem to affect the way the standard protein reads, you have two
alternatives:
1.
Prepare and use a special standard curve to use with samples which are in that
buffer. The standards should be uniformly spiked, say with 100 uL of the
buffer. (Note also that you will have to use a uniform amount of sample+buffer
(100 uL in this example) so that the same amount of the offending interfering
substance is the same in each tube). This isn’t really the best way but is
usually OK and certainly faster than the alternative.
2.
Dialyze or otherwise remove the offending substance from the sample.
In the Lowry protein assay, a common variation (called the “Peterson Modified
Lowry) has you precipitate the tiny amount of protein in the assay with 10%
TCA (Trichloroacetic Acid), and re-dissolve in NaOH before using the Lowry
reagents. This is especially useful when working with proteins that require the
presence of detergents for their integrity (some membrane proteins for
example) because detergents are notorious for interfering with protein assays.
This method allows you to remove the detergent only from that small fraction
which will be consumed anyway by the protein assay
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D. COLUMN SELECTION
Choice of Chromatography media (resin )
The key element to this purification step is the functional group on the resin, DEAE. The
original protocol calls for DEAE Sepharose CL6B (DEAE attached to a Sepharose CL-6B
support). This resin has a recommended flow rate of up to 60 cm/hr. We also have DEAE
Sepharose Fast Flow (recommended working flow rate 100-300 cm/hr). Sepharose CL-6B is
itself a gel filtration resin, but for very large proteins (including multimeric species). Its use in
this protocol is likely historical (that is, the protocol was written before the newer Fast Flow
resins were available).
How do we figure how much resin to use?
This is an ion exchange column, so the goal is to use enough so that all of the -gal will bind
in the lower salt, and then be eluted in the higher salt. However, we have a pretty complex
fraction here, with many proteins other than -gal, and we have no idea whether they will bind
or not. Also, different proteins bind the resin differently, (mg/mL of resin), in a way is
unpredictable (see the empirical Table 10)
I would go with the worst case scenario: let’s say that all of the proteins in the fraction you
plan to load will bind to the column, but may do so poorly. We need to make space for all of
them to bind, ideally in the top 10% of the column.
Lets say we are planning to load 6 mg of protein on the column. If it all binds as
poorly as, say , thyroglobulin, (3 mg thryoglobulin/mL of gel, from Table 10) then we
would need 2 mL of resin, but since we want to use only 1/10 of the resin for that
initial binding, we should aim for a 20 mL bed volume.
Once you have done this calculation for the protein sample you plan to load, check the amount
of resin available, and the column choices.
What size of column should be used?
What do we have in the way of columns (and adaptors) that will work for the amount of resin
we need? Do you want to use a short /fat column, or a tall skinny one? In general, tall skinny
columns give better resolution than short fat ones. But, as you know, skinny columns have to
be run at slower flow rates. Calculate the flow rate for the column you expect to use. How
many minutes will it take to run one column volume through? Consider the volume that you
need to load, and how many will be required to get the A280 back to near-baseline (it won’t be
exactly the same, but check back to your previous ion exchange column). Also remember
that for the gradient, you will need between 5 and 10 column volumes. (The more you have,
the better the resolution, but the longer it will take!)
12
Now, if it looks like your estimates will be practical (i.e., you really don’t want to run a
column that takes three days, and it would not be fair to use more than half of the available
resin!) go ahead. If not, you should consider a smaller or larger column. (You can probably
justify a smaller column because your estimates of how much protein would bind were really
“worst case”. Likely a lot of protein will not bind (especially in 0.2 M NaCl!), and those that
do are likely to bind better than we figured (based on the HSA and a-lactalbumin data in Table
10).
What should the flow rate be?
The appropriate range of flow rates to be used will depend on the particular resin and the
cross sectional area of the column. In addition, consider the following guidelines:

For packing and buffer equilibration: you can use any flow rate up to the maximum for
the resin.

For loading the sample: use a flow rate that is at the slow end of the suggested working
range -- or even slower (based on practical considerations (see below) You want to give
your solutes good opportunity to bind the resin.

Washing unbound protein from the column: at some point after loading it would be OK to
speed the washing up a bit -- once the ionic interactions are established, faster flow rate
should not affect binding and may get species that are weakly interacting off more
quickly.

Elution: good news: even though you have several column volumes with which to elute,
you should increase the flow rate to the top of the working range. If you use a very slow
flow rate, your proteins could get ahead of your salt (especially if the support is also a gel
filtration resin, and are in the void volume) and re-bind further down the column, only to
be eluted again when the salt catches up. This can cause ”tailing”, as some of the proteins
are slightly retarded instead of the whole batch eluting at once.
Column Equilibration:
How close should the conductivity of the incoming buffer and effluent really be? That is
generally an operational question: it depends on what you are doing.
In this procedure, the ionic strength of the starting buffer is already pretty high at 0.2
M NaCl. By starting with this concentration, we have essentially already “salted off”
some of the proteins that would have bound at lower ionic strength. If you start much
below 0.2 M NaCl , you lose that benefit. If you are too high, your -gal may fail to
bind. The gradient will run from 0.2M to 0.4 M NaCl, which is relatively narrow. You
probably want to be within 5% of the designated salt, between 0.19 M and 0.21M.
How do we tell what this is? We have a conductivity meter, but the units are Seimens
(or mS) Use your stock solutions to prepare three solutions (0.19 M. 0.2 M and 0.21
13
M NaCl all in NTM), at least 100 mL each. Read and record the conductivity. What
is the slope of mS vs. M? Now, soon after you begin the equilibration, and then
periodically afterwards, read the conductivity of the effluent, (A word of caution: use
at least 10 mL of effluent in a 50-mL tube, because the probe does not work well
unless there is adequate flow through it.)
Cold
This column should be run in the cold. This is mostly to preserve the activity of the enzyme,
and also keeps the pH correct (since NTM is Tris based, assuming that you made the Tris in
the cold.). This may be difficult given our facilities. You may want to try using cold buffers,
and keeping the column in the cold when not in use. The longer your enzyme is at room
temperature, the more activity it is likely to use, so do what you can to minimize this.
14
IV.
CALCULATIONS –
How the Enzyme and Protein assays tell us about Yield, Specific Activity and
indicate how well purified our protein is.
First determine
Then calculate:
% Yield
=
Enzyme activity:
Units per mL as per handout
Total Enzyme:
Units/mL x Volume = Units
Total enzyme (units) at the step in question x 100%
Total enzyme (units) initial
If you start out a purification with 50 mL of lysate, and reserve 2 mL for later analysis, you
will continue into the purification with 48 mL. Lets say that you carry out an ammonium
sulfate precipitation, and obtain a supernatant (51 mL, increased in volume due to salt), and a
pellet that you resuspend in 10 mL of buffer (resulting in a final measured results
You carry out the enzyme assay on the sup and on the pellet, and from the 2 mL that you had
reserved in the fridge, and get the following results:
Fraction
Volume
in mL
Lysate
AS Sup
AS Pellet
48.0
51.0
11.0
Enzyme
conc
in U/mL
5830
2941
7272
Total
enzyme
In Units
%
Yield
First calculate the number of units of enzyme you have in each fraction, based not the volume
you have of each. NOTE: You use as a starting point the 48 mL that went into the
purification, not the 2 mL that you held aside (you could also call this 50mL – with the 2 mL
that you held out being part of your overall “loss”, but you DO NOT use 2 mL here.
Fraction
Volume
in mL
Lysate
AS Sup
AS Pellet
48.0
51.0
11.0
Enzyme
conc
in U/mL
5830
2941
7272
Total
enzyme
In Units
279,840 U
149,991 U
79,992 U
%
Yield
Protein
conc
mg/mL
Total
Protein
In mg
Specific
Activity
100 %
53 %
29 %
In protein purification, the yield generally refers to the recovery of the protein you are
interested in. Thus it is based on what ever means you are using to detect THAT protein (not
total protein from the protein assay). So here it is based on the enzyme assay, and expressed
as a percentage of what you started with.
In the best possible outcome, your sup and pellet would sum to 100%, but that seldom
happens. Why?
15
Enzyme can be physically lost (as in when you dropped one of the tubes on the floor) or it can
lose its activity (if you boil this enzyme, it will still be there but it will not work). Right now,
you cannot tell how much contribution was one or the other.
Here is another important point – if you are going to carry on with this purification, which
fraction do you want to continue with? If you guessed the AS Pellet because it has a higher
concentration of enzyme that would be WRONG!
It doesn’t much matter what the concentration is. A given amount of enzyme at a high
concentration can be made less concentrated by adding water – and can be made more
concentrated by removing it. We want the greater AMOUNT. In this case, that would be the
supernatant.
But we really would like to know that this is somewhat more pure than what we started with
too. So we look to the protein concentration, which we determine in order to assess the purity
of the enzyme, and also to indicate if we had any significant losses of protein.
Now consider results from the protein assay
Determine
used)
Protein concentration:
Calculate
Total protein :
Fraction
Lysate
AS Sup
AS Pellet
Volum
e
in mL
48.0
51.0
11.0
Enzyme
conc
U/mL
5830
2941
7272
mg/mL (remember to account for vol & dil
mg/mL x vol = mg (fill in the table below)
Total
enzyme
Units
279,840 U
149,991 U
79,992 U
%
Yield
100 %
53 %
29 %
Protein
conc
mg/mL
5.5
1.8
10.9
Total
Protein
mg
Specific
Activity
U/mg
You should calculate the % of protein recovered, but I would not put it in the table, because it
gets confusing. The reason for looking at it is to see how it compares overall with your
enzyme recovery. In this example we recovered about 190 mg in sup + pellet, out of 200 that
we started with, or 95% of the protein we started with (no tube dropped on the floor here!).
However, we only recovered 80% of the enzyme activity. That suggests that some of the
enzyme lost activity. Not good, but good to know!
With the information we have, we can now calculate Specific activity
If you look at the numbers we have so far, you can see that in our pellet we have more protein
that we have in the sup. Because we have more enzyme in the sup, it is great that we have less
protein there – we have gotten most of the enzyme, but lost most of the protein – presumable
other proteins that we do not want. The calculation we use to express this ratio is
Specific Activity:
Units of enzyme
= Units/mg
mg of protein
16
This can be calculated from the total units and mgs, or from the concentrations (because if you
divide units/mL by mg/mL, the mL cancel out and you get Units/mg)
In purification, the idea is to retain as much of my protein (indicated by the activity) and lose
as much irrelevant protein (total mg). The more units per mg, the closer to absolute purity
(i.e., “purified to homogeneity” ) they are. [The maximum specific activity is usually not a
specific number being aimed for – it depends too much on the way the protein of interest is
being measured, and the proportion of the total protein in the initial lysate]
Calculate the specific activity in the table above.
Now you have a stronger argument for keeping the sup – not only is there more of our enzyme
there, but there is also less protein and so the specific activity is higher.
Finally, we can bring in the concept of Fold purification, which is a way to express the
improvement in the ratio of protein if interest relative to all the others at each step of the
purification
Fold purification =
Specific activity of any fraction
Specific activity of initial lysate.
By definition, the “fold purification” of the starting extract = 1. Fill in the table below:
Fraction
Lysate
AS Sup
AS Pellet
Vol
in
mL
48.0
51.0
11.0
Enzyme
conc
U/mL
5830
2941
7272
Total
enzyme
Units
279,840 U
149,991 U
79,992 U
%
Yield
100 %
53 %
29 %
Protein
conc
mg/mL
5.5
1.8
10.9
Total
Protein
mg
264
120
92
Specific
Activity
U/mg
1060
1634
667
Fold
purificatio
n
1
(The reason we do this is to “normalize” the expression of specific activity, so that it does not
depend on the actual units we get – because people often use different assays, and would get
different numbers than cannot be compared. Also, it allows us to conveniently express the
purity, or the success of the purification of different samples.
When describing this, we do not say that the Sup has a “Fold purification of 1.5” , but we say
that the “sup is 1.5-fold purified relative the lysate”. The pellet on the other hand, is less pure.
This can also be used to compare the effectiveness of any step, by substituting the prior
fraction for the initial, to see how effective each step is by itself:
Sp act of column pool II
Sp act of dialyzed ammonium sulfate pellet.
17
This would tell you how much purification you got from the column itself (if you loaded the
column with the pellet sample), which is a good way to assess the performance of any given
step of a purification procedure.
What is a good “fold purification”?
It depends on the system. For any step, a 2-fold purification is not very good, but can be OK
if you are trying to get rid of specific contaminants. On the other hand, some very selective
techniques can give a 100-fold increase in specific activity. The purifications multiply: two
consecutive steps each with a 5-fold purification will give you a product that is 25X more pure
than the starting material.
It also depends on how abundant your protein is. If it is very abundant, say 5 % by weight of
the total protein, its maximum purification would be 20-fold.
if you start with a lysate containing 40 mg of protein and 100 units of enzyme, but by
weight the lysate was 5 % of your enzyme, that would be 2 mg, so the best you could
do in purification would be 100 units/2 mg, or 50 U/mg. Compared to the original
lysate (100 units/40 mg, or 2.5 U/mg, purification achieved would be 20-fold (50/2.5)
Final assays require tables of exactly the same design as the first ones (see CALCUATIONS
SECTIONS) , but this will have additional samples:
For the Purification table:
Fraction
Vol
mL
Enz
conc
U/mL
Total
enzyme
Units
%
Yield
Lysate
AS Pellet
AS Pellet, dialyzed
Unbound
Pool I
Pool II
Pool III
More if needed
18
Protein
conc
mg/mL
Total
Protein
mg
Specific
Activity
U/mg
Fold
purific