10 Things
Every
Molecular
Biologist
Should Know
a BitesizeBio.com eBook
Nick Oswald
Suzanne Kennedy
Megan Hogan
Megan Cartwright
Edited By Nick Oswald
Preface
Molecular Biology is a funny old business...
Preface
Molecular biology is a funny old
business. The ton of theoretical
knowledge that we cram in during our
undergrad years is scant preparation
for life at the bench and leaves a lot to
be said.
When we first hit the lab there are so
many things to learn before we even
get started that many things go
unlearned.
How specific techniques work, what
actually is in those kits we use and why
we use certain approaches rather than
others.
Often these nuggets of knowledge can
mean the difference between
understanding where an experiment is
going wrong and not understanding,
between getting a result and not getting
one.
But normally, these nuggets can’t be
found in lectures or in text books.
Instead they must be gleaned through
hard on-the-job experience or passed
down by word-of-mouth from more
experienced mentors.
One of the core aims of Bitesize Bio
(www.bitesizebio.com) is to make it
easier to learn on the job as a molecular
biologist by providing a place where
these nuggets of vital, often over-looked
knowledge can be found.
This short eBook provides 10 such
nuggets, neatly contained in one place.
From How SDS-PAGE works to the
chemical reason why enzymes have
optimal temperatures we hope that there
will be something in here that will
enlighten and entertain even the most
experienced scientist.
If you like the information you find in this
eBook - which is the first of many we’ll
be creating on a variety of topics - we
would love you pass the file onto your
friends to help spread the word about
what we are doing at Bitesize Bio.
And if you are hungry for more please be
sure to visit us at BitesizeBio.com to get
the latest articles and eBooks we have
on offer.
Finally if you have any suggestions for
articles you’d like to see on Bitesize Bio,
if you’d like to write something for the
website yourself, or if you’d just like to let
us know what you think (good or bad!) of
our efforts, please feel free to get in touch
via the contact page at BitesizeBio.com.
Thanks for reading.
The Bitesize Bio team.
ii
How SDS-PAGE Works
“If you are still wondering why the stacking gel is needed, think of
what would happen if you didn’t use one.”
Section 01 By Nick Oswald
SDS-PAGE (sodium dodecyl sulphatepolyacrylamide
gel electrophoresis) is
commonly used in the lab for the
separation of proteins based on their
molecular weight. It’s one of those
techniques that is commonly used but
not frequently fully understood. So let’s
try and fix that.
SDS-PAGE separates proteins
according to their molecular weight,
based on their differential rates of
migration through a sieving matrix (a
gel) under the influence of an applied
electrical field.
Making the rate of protein
migration proportional to
molecular weight.
The movement of any charged species
through an electric field is determined
by it’s net charge, it’s molecular radius
and the magnitude of the applied field.
But the problem with natively folded
proteins is that neither their net charge
nor their molecular radius are molecular
weight dependent. Instead, their net
charge is determined by amino acid
composition i.e. the sum of the positive
and negative amino acids in the protein
and molecular radius by the protein’s
tertiary structure.
So in their native state, different proteins
with the same molecular weight would
migrate at different speeds in an
electrical field depending on their charge
and 3D shape.
To separate proteins in an electrical field
based on their molecular weight, we
need to destroy the tertiary structure by
reducing the protein to a linear molecule,
and somehow mask the intrinsic net
charge of the protein. That’s where SDS
comes in.
The Role of SDS (et al)
SDS is a detergent that is present in the
SDS-PAGE sample buffer where, along
with a bit of boiling, and a reducing agent
(normally DTT or B-ME to break down
protein-protein disulphide bonds), it
disrupts the tertiary structure of proteins.
This brings the folded proteins down to
linear molecules.
iv
SDS helps to disrupt the tertiary
protein structure
SDS also coats the protein with a
uniform negative charge, which masks
the intrinsic charges on the R-groups.
SDS binds fairly uniformly to the linear
proteins (around 1.4g SDS/ 1g protein),
meaning that the charge of the protein
is now approximately proportional to it’s
molecular weight.
SDS is also present in the gel to make
sure that once the proteins are
linearized and their charges masked,
they stay that way throughout the run.
The dominant factor in determining the
migration rate of an SDS-coated protein
in a gel is it’s molecular radius.
SDS-coated proteins have been shown
to be linear molecules, 18 Angstroms
wide and with length proportional to
their molecular weight, so the molecular
radius (and hence their mobility in the
gel) is determined by the molecular
weight of the protein. Since the SDScoated
proteins have the same charge
to mass ratio there will be no differential
migration based on charge.



The gel matrix

In an applied electrical field, the SDStreated
proteins will now move toward
the cathode at different rates depending
on their molecular weight. These
different mobilities will be exaggerated
by the high-friction environment of a gel
matrix.
As the name suggests, the gel matrix
used for SDS-PAGE is polyacrylamide,
which is a good choice because it is
chemically inert and, crucially, can easily
be made up at a variety of concentrations
to produce different pore sizes which, in
turn, can produce varying separating
conditions that can be adjusted to suit your
needs.
The discontinuous buffer
system and the stacking gel
To conduct the current through from the
anode to the cathode through the gel, a
buffer is obviously needed. Mostly we use
the discontinuous Laemmli buffer system.
“Discontinuous” simply means that the
buffer in the gel and the tank are different.
Typically, the system is set up with a
stacking gel at pH 6.8, buffered by TrisHCl,
a running gel buffered to pH 8.8 by
Tris-HCl and an electrode buffer at pH 8.3.
The stacking gel has a low concentration of
acrylamide and the running gel has a
higher concentration capable of retarding
the movement of the proteins.
So what’s with all of those different pH’s?
Well, glycine can exist in three different
charge states, positive, neutral or negative
v
The discontinuous buffer system
depending on the pH. This is shown in
the diagram below. Control of the
charge state of glycine by the different
buffers is the key to the whole stacking
gel thing.
So here’s how the stacking gel works.
When the power is turned on, the
negatively-charged glycine ions in the
pH 8.3 electrode buffer are forced to
enter the stacking gel, where the pH is
6.8. In this environment glycine
switches predominantly to the
zwitterionic (neutrally charged) state.
This loss of charge causes them to
move very slowly in the electric field.
The Cl- ions (from Tris-HCl) on the
other hand, move much more quickly in
the electric field and they form an ion
front that migrates ahead of the glycine.
The separation of Cl- from the Tris
counter-ion (which is now moving
towards the cathode) creates a narrow
zone with a steep voltage gradient that
pulls the glycine along behind it,
resulting in two narrowly separated
fronts of migrating ions; the highly
mobile Cl- front, followed by the slower,
mostly neutral glycine front.
All of the proteins in the gel sample have
an electrophoretic mobility that is
intermediate between the extreme of the
mobility of the glycine and Cl- so when
the two fronts sweep through the
sample-well the proteins are
concentrated into the narrow zone
between the Cl- and glycine fronts.
And they’re off!
This procession carries on until it hits the
running gel, where the pH switches to
8.8. At this pH the glycine molecules are
mostly negatively charged and can
migrate much faster than the proteins.
So the glycine front accelerates past the
proteins, leaving them in the dust.
The result is that the proteins are dumped
in a very narrow band at the interface of the
stacking and running gels and since the
running gel has an increased acrylamide
concentration, which slows the movement
of the proteins according to their size, the
separation begins.
What was all of that about?
If you are still wondering why the stacking
gel is needed, think of what would happen
if you didn’t use one.
Gel wells are around 1cm deep and you
generally need to substantially fill them to
get enough protein onto the gel. So in the
absence of a stacking gel, your sample
would sit on top of the running gel, as a
band of up to 1cm deep.
Rather than being lined up together and
hitting the running gel together, this would
mean that the proteins in your sample
would all enter the running gel at different
times, resulting in very smeared bands.
vi
The effect of buffer pH on glycine
charge
So the stacking gel ensures that all of
the proteins arrive at the running gel at
the same time so proteins of the same
molecular weight will migrate as tight
bands.
Separation
Once the proteins are in the running
gel, they are separated because higher
molecular weight proteins move more
slowly through the porous acrylamide
gel than lower molecular weight
proteins. The size of the pores in the
gel can be altered depending on the
size of the proteins you want to
separate by changing the acrylamide
concentration.
For a broader separation range, or for
proteins that are hard to separate, a
gradient gel, which has layers of
increasing acrylamide concentration,
can be used.
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vii
How Proteases and
Protease
Inhibitors Work
“To defeat the protease, you must become the protease”
Section 02 By Megan Cartwright
Proteases: wild, mysterious,
destructive.  What are these untamed
elements ravaging your precious
lysate?
How can a drop of EDTA or a smidge of
“cocktail” protect that sample, which is
gently cradling your hopes, your
dreams, and your desire to survive the
next lab meeting?
Brace yourself for a biochem flashback:
in this section, I’ll explain the what and
how of proteases and protease
inhibitors, and we’ll do it with phrases
like “reversible competitive inhibition”
and “zymogen.”
Proteases: the good, the
bad, the hydrolytic
All organisms have proteolytic and
phosphorylating enzymes.  You.  Me. 
Helicoverpa armigera.  While involved
in everything from host defense to
wound healing to viral replication,
proteases can be broadly lumped into
two camps based on what’s in their
active site and how it interacts with the
target peptide bond. 
Camp One hydrolyzes the bond through
nucleophilic activation of a serine,
threonine, or cysteine at its active site. 
Camp Two, which includes the aspartic,
glutamic, and metalloproteases, uses
water itself for the hydrolysis.
These tidy camps break down under
questioning about preferred targets and
substrate affinity.  Although Camp One’s
serine proteases all agree that an active
site serine is superior to the plebeian
cysteine, they still have tense family
dinners because of each of the
proteases has a different target
preference and structure.
These structural differences drive a
protease’s specificity for where it will
attack: terminal hydrolysis for the
exopeptidases, internal hydrolysis for the
endopeptidases. 
Although the exopeptidases only choose
between the amino or carboxyl terminus,
the endopeptidases are gloriously picky. 
To go back to the serine example, our
friend trypsin hydrolyzes on the carboxyl
side of positively charged arginine or
lysine, unless these are followed by a
proline. 
And while its buddy chymotrypsin is also a
serine protease, chymotrypsin simply
doesn’t touch arginine or lysine and really
just prefers aromatics like tyrosine,
tryptophan, or phenylalanine.
The last thing a cell wants is these
enzymes running amok, rampaging willynilly
through the cytoplasm, chewing offtarget,
and generally behaving like – I say
this with great familiarity – Ohio State
football fans. 
Regulation and segregation are the cell’s
prime approaches for dealing with these
enzymes: regulation by, for example,
holding them in an inactive precursor state
(a “zymogen”) and segregation to the
lysosome or secretory pathways, among
other locations.
ix
Inhibitors: You can’t stop
with just one
All that chemical lysis, scraping,
sonication, and freeze-thawing you
inflict throws the whole system into
chaos.  No peptide bond is safe! 
Exopeptidases attack the ends of
amino acid chains while
endopeptidases hydrolyze their internal
links.  Kinases phosphorylate and
dephosphorylate at will.  Good luck
quantifying NOS induction or making a
case for p53 activation by Western. 
Without inhibitors, your results will be
inconsistent, your lab meetings
agonizing, and your graduation date a
pipe dream.
Tragically, there is no Wunderkind
inhibitor that takes out all proteases,
halts every phosphorylation event, and
does some light tidying-up about the
centrifuges.  It takes a “cocktail” of
approaches to stave off proteolysis. 
Protease inhibitors can act irreversibly,
reversibly, and reversibly-under-duress,
and can be anything from small
molecules like sodium fluoride to big
honkin’ polypeptide analogues of
evolution’s own microbial inhibitors. 
They can compete with substrates for an
active site – sometimes destroying the
site in the process – or selfishly hoard
the precious, precious cations needed
for protease function.
Competitive inhibitors act by binding in a
substrate-like manner to the active site to
block it or by binding and destroying it
through modifications such as
esterification.  Many large competitive
inhibitors boost their affinity and
specificity by binding to the overlooked
wallflower regions neighboring the hogsall-the-glory
active site.  You have likely
already come across competitive
inhibitors.  They include all sorts, from
aprotinin, a 6 kDa polypeptide serine
protease that is reversible if you ask
politely at pH extremes, to sodium
orthovanadate, a small molecule which
inhibits the ever-gullible protein
phosphotyrosyl phosphatases.
By contrast, the toddler-like chelators
(motto: MINE!) take advantage of their
enhanced affinity for metal ions to complex
with them so that nobody else can react
with them.  This matters because
metalloproteases need zinc, magnesium,
manganese, calcium, or even cobalt in their
active site to activate water to hydrolyze
peptide bonds to meet their quarterly
hydrolysis goals.  Take away that zinc with
EDTA, complex up that calcium with EGTA,
and it’s bye-bye metalloprotease activity.
Conclusion: To defeat the
protease, you must become
the protease
Proteases are just some of the destructive
forces nature aims at your beloved cell
lysates, and protease inhibitors are one of
the ways that the clever scientist can, judolike,
turn the universe against itself.  For
more information on beating the universe
into submission, check out Emily Crow’s
“How To Preserve Your Samples In
Western Blotting”, Jode Plank’s “Streamline
Your Western Blots”, on Bitesize Bio.
 
x
Agarose Gels Do Not
Polymerize
“Polyacrylamide gel polymerization - Try saying that three times
quickly!”
Section 03 By Nick Oswald
I was browsing a certain website the
other day when I came across a
protocol that advised: “When agarose
starts to cool, it undergoes what is
known as polymerization”
Polymerizing agarose? Someone is
getting their gel matrices mixed up, and
it’s not the first time I have heard this
said, so it looks like some myth-busting
is required.
So hunker down for a quick run down
on the difference between polymerizing
and non-polymerizing gel matrices.
Of the common gel matrices used in
molecular biology, polyacrylamide, agar
and agarose, polyacrylamide is the one
that polymerizes. I’ll deal with agarose
and agar later.
Polyacrylamide gel
polymerization
Polyacrylamide, used mainly for SDSPAGE,
is a matrix formed from
monomers of acrylamide and bisacrylamide.
It’s strengths are that is it
chemically inert – so won’t interact with
proteins as they pass through – and that
it can be easily and reproducibly made
with different pore sizes to produce gels
with different separation properties.
The polymerization reaction, shown in
the diagram below, is a vinyl addition
catalyzed by free radicals. The reaction
is initiated by TEMED, which induces
free radical formation from ammonium
persulphate (APS). The free radicals
transfer electrons to the acrylamide/
bisacrylamide monomers, radicalizing
them and causing them to react with each
other to form the polyacrylamide chain.
In the absence of bis-acrylamide, the
acrylamide would polymerize into long
strands, not a porous gel. But as the
diagram shows, bis-acrylamide cross-links
the acrylamide chains and this is what
gives rise to the formation of the porous gel
matrix. The amount of cross-linking, and
therefore the pore size and consequent
separation properties of the gel can be
controlled by varying the ratio of acrylamide
to bis-acrylamide.
xii
Polyacrylamide gel polymerization - Try saying that three times quickly!
For more information on polyacrylamide
gel polymerization see Biorad Bulletin
1156
Agarose gel formation
So what about agarose? Well, agarose
– the main component of the gelatinous
agar that can be isolated from certain
species of seaweed – is itself a
polymer. But, polymerization is not the
mechanism for agarose gel formation.
Chemically, agarose is a
polysaccharide, whose monomeric unit
is a disaccharide of D-galactose and
3,6-anhydro-L-galactopyranose which
is shown in the diagram below.
In aqueous solutions below 35°C these
polymer strands are held together in a
porous gel structure by non-covalent
interactions like hydrogen bonds and
electrostatic interactions. Heating the
solution breaks these non-covalent
interactions and separates the strands.
Then as the solution cools, these noncovalent
interactions are re-established
and the gel forms.
So agarose (and agar) gels form by
gellation through hydrogen bonding
and electrostatic interactions, not
through polymerization.
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xiii
Chemically, agarose is a
polysaccharide, whose
monomeric unit is a disaccharide
of D-galactose and 3,6-anhydro-L-
galactopyranose
How Ethanol Precipitation of
DNA and RNA
Works
“First we need to know why nucleic acids are soluble in water...”
Section 04 By Nick Oswald
Ethanol precipitation is a commonly
used technique for concentrating and
de-salting nucleic acid (DNA or RNA)
preparations in aqueous solution. The
basic procedure is that salt and ethanol
are added to the aqueous solution,
which forces the nucleic acid to
precipitate out of solution.
The precipitated nucleic acid can then
be separated from the rest of the
solution by centrifugation. The pellet is
washed in cold 70% ethanol then after
a further centrifugation step the ethanol
is removed, and the nucleic acid pellet
is allowed to dry before being
resuspended in clean aqueous buffer.
So how does this work?
A bit about solubility…
First we need to know why nucleic
acids are soluble in water. Water is a
polar molecule – it has a partial
negative charge near the oxygen atom
due the unshared pairs of electrons,
and partial positive charges near the
hydrogen atoms (see the diagram
below).
Because of these charges, polar
molecules, like DNA or RNA, can interact
electrostatically with the water
molecules, allowing them to easily
dissolve in water.
Polar molecules can therefore be
described as hydrophilic and non-polar
molecules, which can’t easily interact with
water molecules, are hydrophobic.
Nucleic acids are hydrophilic due to the
negatively charged phosphate (PO3-)
groups along the sugar phosphate
backbone.
The role of the salt…
Ok, so back to the protocol. The role of the
salt in the protocol is to neutralize the
charges on the sugar phosphate backbone.
A commonly used salt is sodium acetate. In
solution, sodium acetate breaks up into Na
+ and [CH3COO]-.
The positively charged sodium ions
neutralize the negative charge on the PO3groups
on the nucleic acids, making the
molecule far less hydrophilic, and therefore
much less soluble in water.
The role of the ethanol…
The electrostatic attraction between the
Na+ ions in solution and the PO3- ions are
dictated by Coulomb’s Law, which is
xv
Polarity of water molecules. Oxygen
atoms are slightly negatively charged,
hydrogens are slightly positive
affected by the dielectric constant of the
solution.
Water has a high dielectric constant,
which makes it fairly difficult for the Na+
and PO3- to come together.
Ethanol on the other hand has a much
lower dielectric constant, making it
much easier for Na+ to interact with the
PO3-, shield it’s charge and make the
nucleic acid less hydrophilic, causing it
to drop out of solution.
The role of temperature…
Incubation of the nucleic acid/salt/
ethanol mixture at low temperatures
(e.g. -20 or -80C) is commonly cited in
protocols as necessary in protocols.
However, according to Maniatis et al
(Molecular Cloning, A Laboratory
Manual 2nd Edition… 2nd edition?? – I
need to get a newer version!), this is
not required, as nucleic acids at
concentrations as low as 20ng/mL will
precipitate at 0-4C so incubation for
15-30 minutes on ice is sufficient.
The wash step with 70%
ethanol…
This step is to wash any residual salt
away from the pelleted DNA.
A few tips on nucleic acid
precipitation…
Choice of salt
▪ Use Sodium acetate (0.3M final conc,
pH 5.2) for routine DNA precipitations
▪ Use Sodium chloride (0.2M final conc)
for DNA samples containing SDS since
NaCl keeps SDS soluble in 70% ethanol
so it won’t precipitate with the DNA.
▪ Use Lithium Chloride (0.8M final conc)
for RNA. This is because 2.5-3 volumes
of ethanol should be used for RNA
precipitation and LiCl is more soluble in
ethanol than NaAc so will not precipitate,
but beware – chloride ions will inhibit
protein synthesis and DNA polymerase
so LiCl is no good for RNA preps for in
vitro translation or reverse transcription. In
these cases, use NaAc.
▪ Use Ammonium acetate (2M final conc)
for the removal of dNTPs, but do not use
for preparation of DNA for T4
polynucleotide kinase reactions as
ammonium ions inhibit the enzyme.
To increase the yield in precipitations of
low concentration or small nucleic acid
pieces (< 100 nucleotides)
▪ Add MgCl2 to a final concentration of
0.01M
▪ Increase the time of incubation on ice
before centrifugation to 1 hour.
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xvi
How Alkaline Lysis Works
“The easiest way to describe how alkaline lysis works is to go
through the procedure and explain each step, so here goes....”
Section 05 By Nick Oswald
Alkaline lysis was first described by
Birnboim and Doly in 1979 (Nucleic
Acids Res. 7, 1513-1523) and has, with
a few modifications, been the preferred
method for plasmid DNA extraction
from bacteria ever since. The easiest
way to describe how alkaline lysis
works is to go through the procedure
and explain each step, so here goes.
1. Cell Growth and
Harvesting
The procedure starts with the growth of
the bacterial cell culture harboring your
plasmid. When sufficient growth has
been achieved, the cells are pelleted by
centrifugation to remove them from the
growth medium.
2. Re-suspension
The pellet is then re-suspended in a
solution (normally called solution 1, or
similar in the kits) containing Tris,
EDTA, glucose and RNase A. Divalent
cations (Mg2+, Ca2+) are essential for
DNase activity and the integrity of the
bacterial cell wall. EDTA chelates
divalent cations in the solution
preventing DNases from damaging the
plasmid and also helps by destabilizing
the cell wall. Glucose maintains the
osmotic pressure so the cells don’t burst
and RNase A is included to degrade
cellular RNA when the cells are lysed.
3. Lysis
The lysis buffer (aka solution 2) contains
sodium hydroxide (NaOH) and the
detergent Sodium Dodecyl (lauryl)
Sulfate (SDS).
SDS is there to solubilize the cell
membrane. NaOH helps to break down
the cell wall, but more importantly it
disrupts the hydrogen bonding between
the DNA bases, converting the doublestranded
DNA (dsDNA) in the cell,
including the genomic DNA (gDNA) and
your plasmid, to single stranded DNA
(ssDNA).
This process is called denaturation and
is central part of the procedure (which is
why it’s called alkaline lysis!). SDS also
denatures most of the proteins in the cells,
which helps with the separation of the
proteins from the plasmid later in the
process.
It is important during this step to make sure
that the re-suspension and lysis buffers are
well mixed, although not too vigorously.
Also remember that SDS and NaOH are
pretty nasty so it’s advisable to wear gloves
and eye protection when performing
alkaline lysis.
4. Neutralization
Addition of potassium acetate (solution 3)
increases the pH of the mixture. Under
these conditions the hydrogen bonding
between the bases of the single stranded
DNA can be re-established, so the ssDNA
can re-nature to dsDNA.
This is the selective part. While it is easy
for the the small circular plasmid DNA to renature
it is impossible to properly anneal
those huge gDNA stretches.
This is why it’s important to be gentle
during the lysis step because vigorous
xviii
mixing or vortexing will shear the gDNA
producing shorter stretches that can reanneal
and contaminate your plasmid
prep.
While the double-stranded plasmid can
dissolve easily in solution, the single
stranded genomic DNA, the SDS and
the denatured cellular proteins stick
together through hydrophobic
interactions to form a white precipitate.
The precipitate can easily be separated
from the plasmid DNA solution by
centrifugation.
5. Cleaning and
concentration
Now your plasmid DNA has been
separated from the majority of the cell
debris but is in a solution containing
lots of salt, EDTA, RNase and residual
cellular proteins and debris, so it’s not
much use for downstream applications.
The next step is to clean up the solution
and concentrate the plasmid DNA.
There are several ways to do this
including phenol/chloroform extraction
followed by ethanol precipitation and
affinity chromotography-based methods
using a support that preferentially binds
to the plasmid DNA under certain
conditions of salt or pH, but releases it
under other conditions. The most
common methods are detailed in the
Bitesize Bio article on 5 ways to clean up
a DNA sample.
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xix
How Silica Spin Column
DNA and RNA
Preps Work
“...the downside [of using kits] is that we don't actually know what is in the
mysterious... set of solutions which... makes troubleshooting more difficult.”
Section 06 By Suzanne Kennedy
These days, most labs use commercial
kits, which employ spin columns, for the
isolation of nucleic acids. The spin
columns contain a silica resin that
selectively binds DNA/RNA, depending
on the salt conditions and other factors
influenced by the extraction method.
When things are going well, these kits
make the whole process much easier
and faster than the 'Methods of Old'.
However, the downside is that we don't
actually know what is in the mysterious
and proprietary set of solutions which,
of course, makes troubleshooting more
difficult.
In this section I’ll explain how silica spin
filter kits work and what is going on at
each step.
I’ll also go over some common
problems specific to silica columns that
can be overcome or avoided with just a
little extra understanding.
Lysis
The lysis approach may vary based on
the whether you want DNA or RNA, but
the common denominator is a lysis buffer
containing a high concentration of
chaotropic salt.
Chaotropes destabilize hydrogen bonds,
van der Waals forces and hydrophobic
interactions. Proteins are destabilized,
including nucleases, and the association
of nucleic acids with water is disrupted
setting up the conditions for their transfer
to silica.
Chaotropic salts include guanidine HCL,
guanidine thiocyanate, urea, and lithium
perchlorate.
Besides the chaotropes, lysis buffer
usually contains detergents to help with
protein solubilization and lysis.
There can also be enzymes used for
lysis depending on the sample type.
Proteinase K is one of these, and
actually works very well in these
denaturing buffers; the more denatured
the protein, the better Proteinase K works.
Lysozyme, however, does not work in the
denaturing conditions of the lysis buffer, so
lysozyme treatment is usually done before
adding the denaturing salts.
For plasmid preps, the lysis is very different
from extraction for RNA or genomic DNA
because the plasmid has to be separated
from the genomic DNA first.
If you throw in chaotropes, you’ll release
everything at once and won’t be able to
differentially separate the small circular
DNA from the high molecular
weight chromosome.
So in plasmid preps the chaotropes are not
added until after lysis and the salts are
used for binding.
A more in-depth explanation of alkaline
lysis for plasmid preps can be found in the
previous section “How Alkaline Lysis
Works”
xxi
Binding
The chaotropic salts are critical for
lysis, but also for binding of the nucleic
acids to the silica resin.
To enhance the binding of nucleic acids
to silica, alcohol is also added at the
binding stage. Most of the time this is
ethanol but sometimes it may be
isopropanol.
The concentration of ethanol, and
the volume used during binding, both
have big effects. Too much
(concentration or volume) and you’ll
bring in a lot of degraded nucleic acids
and small species that will influence
UV260 readings and throw off some
of your yields.  Too little, and it may
become difficult to wash away all of the
salt from the membrane.
The important point here is that the
ethanol influences binding and the
amount added is optimized for
whatever kit you are using. Modifying
that step can help change what you
recover so if you are having problems
and want to troubleshoot recovery, this
can be a step to evaluate further.
Another way to diagnose problems is to
save the flow-through after binding and
precipitate it to see if you can find the
nucleic acids you are searching for. If
you used an SDS-containing detergent in
lysis, try using NaCl as a precipitant to
avoid contamination of the DNA or RNA
with detergent.
Washing Steps
Your lysate was centrifuged through the
silica membrane and now your DNA or
RNA should be bound to the column and
the impurities, protein and
polysaccharides, should have passed
through. 
But, the membrane is still dirty
with residual proteins and salt. If the
sample was from plants, there will still be
polysaccharides, maybe some pigments
too, left on the membrane, or if the
sample was blood, the membrane might
be tinted brown or yellow.
The wash steps serve to remove these
impurities.  There are typically two washes,
although this can vary depending on the
sample type. The first wash will often have
a low amount of chaotropic salt to remove
the protein and colored contaminants. This
is always followed with an ethanol wash to
remove the salts. If the prep is something
that didn’t have a lot of protein to begin
with, such as plasmid preps or PCR clean
up, then only an ethanol wash is needed.
Removal of the chaotropic salts is crucial to
getting high yields and purity of DNA or
RNA. Some kits will even recommend
washing the column twice with ethanol. If
salt remains behind, the elution of nucleic
acid is going to be poor, and the A230
reading will be high, resulting in low
260/230 ratios.
Dry Spin
After the ethanol wash, most protocols
have a centrifugation step to dry the
column. This is to remove the ethanol and
is essential for a clean eluant. When 10
mM Tris buffer or water is applied to the
xxii
membrane for elution, the nucleic acids
can become hydrated and will release
from the membrane. If the column still
has ethanol on it, then the nucleic acids
cannot be fully rehydrated.
Skipping the drying step results in
ethanol contamination and low yields.
The main indication of this problem
comes when you try to load the sample
onto an agarose gel; the DNA will not
sink, but floats in the buffer, even in the
presence of loading dye.  Another
indicator of ethanol contamination is
that if you put the sample in the -20C, it
doesn’t freeze.
Elution
The final step is the release of pure
DNA or RNA from the silica. For DNA
preps, 10 mM Tris at a pH between 8-9
is typically used. DNA is more stable at
a slightly basic pH and will dissolve
faster in a buffer. This is true even for
DNA pellets. Water tends to have a low
pH, as low as 4-5 and high molecular
weight DNA may not completely
rehydrate in the short time used for
elution. Elution of DNA can be
maximized by allowing the buffer to sit in
the membrane for a few minutes before
centrifugation.
RNA, on the other hand, is fine at a
slightly acidic pH and so water is the
preferred diluent. RNA dissolves readily
in water.
What other things can go
wrong with silica spin
columns?
Low yields: If you experience yields
lower than you expected for a sample,
there are many factors to think about.
Usually it is a lysis problem, and
incomplete lysis is a very common cause
of low yields.
Alternatively, the problem could be
incorrect binding conditions. 
Make sure to use fresh high quality ethanol
(100% 200 proof) to dilute buffers or for
adding to the binding step.
Low quality ethanol or old stocks may have
taken on water and not be the correct
concentration. If the wash buffer is not
made correctly, you may be washing off
your DNA or RNA.
Low Purity: If the sample is contaminated
with protein (low 260/280) then maybe you
started with too much sample and the
protein was not completely removed or
dissolved. 
If the sample has poor 260/230 ratio the
issue is usually salt from the bind or the
wash buffer. Make sure that the highest
quality ethanol was used to prepare wash
buffers and if the problem continues, give
the column an additional wash.
Some samples have a lot more inhibitors
compared to others. Environmental
samples are especially prone to purity
issues because humic substances are
solubilized during extraction. Humics
behave similarly to DNA and are difficult to
remove from the silica column. For this
type of sample, specialized techniques
xxiii
exist to remove the protein and humics
prior to the column step.
Degradation: This is more of a
concern for RNA preps and an article
that gives specific advice can be found
in the Bitesize Bio article
“Troubleshooting RNA Isolation”. 
Mainly with RNA, degradation occurs
from improper storage of the sample or
an inefficient lysis, assuming of course
that you eluted with RNase-free water.
For DNA preps, degradation is not a
huge problem because for PCR, the
DNA can be sheared and it works fine.
But if you were hoping not to have so
much sheared DNA, then you may
have used too strong a lysis method.
PCR Clean-up Special
Considerations: PCR clean-up is the
easiest of all silica spin column
techniques, because it simply involves
adding a high concentration of binding
salts (typically between 3-5 volumes of
salt per volumes of PCR reaction) and
centrifuging the column.
So when PCR Clean-up kits fail, it can
be particularly frustrating.  The first
question I ask people is “did you check
the results of the PCR on a gel?”
because you cannot UV check a PCR
reaction and get an accurate reading.
There is way too much in a PCR reaction
absorbing UV at 260: nucleotides,
detergents, salts, and primers. In my
experience, apparent failure of a PCR
clean-up kit to work frequently is
normally caused by a PCR reaction that
has failed and so there was nothing to
clean up in the first place.
But if you know you had a strong PCR
product, the best approach is to just save
your flow-through fraction after binding. If
the DNA doesn’t bind, that’s where it is.
You can always rescue it and then clean
it up again. And then call tech support
and ask for a replacement kit.
If you liked this, you’ll
also like:
Recycle Those DNA Extraction
Columns
by Liam Thompson at BitesizeBio.com
You know those ridiculously priced and
throw-away DNA mini, midi and maxi-prep
columns? Well the good news is that you
can actually re-use them if you are
reasonably careful at regenerating them,
with this simple and cheap method..
...[click to read more]
xxiv
How DNA Ligation Works
“Don’t be scared off by the chemistry, this is worth taking some
time to understand!”
Section 07 By Nick Oswald
DNA ligase (EC 6.5.1.1) covalently
joins the phosphate backbone of DNA
with blunt or compatible cohesive ends
(see figure below) and it’s natural role
is in repairing double strand breaks in
DNA molecules.
In molecular biology it is commonly
used for the insertion of restriction
enzyme-generated DNA fragments into
vector backbones. Commercial ligases
are supplied with a reaction buffer
containing ATP and Mg2+, which are
both essential for ligase activity.
Since ATP can be damaged by
repeated freeze-thaw cycles, it is
advisable to make aliquots of the buffer
(see the Bitesize Bio article “5 DNA
ligation tips” for more information).
The ligation reaction itself has two
basic steps. Firstly the DNA ends have
to collide by chance and stay together
long enough for the ligase to join them.
This is the most inefficient part of the
reaction, but is easier at low
temperatures.
Why? Well, as you will probably know, all
molecules move faster at higher
temperatures so you can imagine that it
is going to be easier for two DNA ends to
collide and stay together if they are
gently floating through the solution at low
temperature, rather than whizzing about
as they would be at higher temperatures.
For cohesive ends, there is an additional
reason; lower temperatures stabilize the
hydrogen bonding between the
complementary nucleotides, which really
helps to keep things in place.
The second step is the enzymatic reaction,
which is shown schematically on the next
page.
DNA ligase catalyzes the joining of the 3′OH
to the 5′-phosphate via a two step
mechanism. First, the AMP nucleotide,
which is attached to a lysine residue in the
enzyme’s active site, is transferred to the
5′-phosphate. Then the AMP-phosphate
bond is attacked by the 3′-OH, forming the
covalent bond and releasing AMP.
To allow the enzyme to carry out further
reactions the AMP in the enzyme’s active
xxvi
The DNA ligase can covalently join blunt ends, but cohesive ends are a bit easier
because the hydrogen bonds hold the cohesive ends in the right place for the
enzyme to do the job.
site must be replenished by ATP.
The DNA ligase enzyme has optimal
activity at 25°C so the ligation reaction
is carried out at a temperature that is a
trade-off between the optimal
temperatures for bringing the DNA ends
together (1°C) and the enzymatic
reaction (25°C). Normally 1hr at 16°C is
fine but since bringing the DNA ends
together is the least efficient part of the
reaction favoring this by lowering the
temperature to 4°C can give even more
efficiency.
However, the enzyme will work very
slowly at this temperature so a long
(e.g. overnight) incubation time is
required.
xxvii
The DNA ligation mechanism. Don’t be scared off by the chemistry, this is worth
taking some time to understand!
If you liked this, you’ll also like:
Pinpointing DNA Ligation Problems
by Nick Oswald at BitesizeBio.com
How Colorimetric Assays
Work
“...even the most traditional of molecular biologists perform
chemical reactions nearly every day”
Section 08 By Megan Hogan
One of my colleagues, a very good
molecular biologist, told me that the
only time she uses chemistry is when
she needs to calculate molarities.
I, of course, scoffed at this statement,
and tried to remind her of all the
chemistry she uses daily.
I may be a bit biased since I am a
chemist, but surely, all the
chromophores, fluorophores, and
imaging probes, etc., involve some
knowledge of chemistry and chemical
reactions. She shrugged, and replied
that they’re all supplied in kits.
So, in defense of all chemistry nerds
out there, here is my rebuttal: even the
most traditional of molecular biologists
perform chemical reactions nearly
every day.
From purification by ion exchange
resins and affinity columns, to
calculating protein concentrations and
coupling bioconjugates, chemistry is
behind it all. For this article I will focus
on a couple of colorimetric assays and
hopefully convince you that knowledge
of these chemical reactions is helpful in
performing these assays and
understanding the results, as well as in
troubleshooting when things go awry.
By taking advantage of the electronic
properties and specific wavelengths of
chromophores, we can deduce a great
deal of information.  Colorimetric assays
help us to distinguish a particular
enzyme from another, to quantitate
catalytic activity, as well as inhibition of
this activity, to generate proliferative and
toxicity profiles, and even to determine
the concentration of proteins in solution.
They are a simple and convenient way to
visualize biological processes. As a
former PI of mine used to say, in his thick
German accent, while pointing to his
eyes, “Nature has provided us with our
own spectrophotometers, no?”
Even more appealing, many colorimetric
assays are commercially available as
kits, usually with a detailed protocol. In
fact, colorimetric assays are used so
regularly in chemistry and biology, that to
cover them all would take an entire
book.  So, for simplicity’s sake, this
article will focus on two phosphatase
assays.
Kinases and phosphatases mediate a
variety of cellular processes like
metabolism, gene transcription and
translation, protein-protein interactions, and
apoptosis, through protein phosphorylation
and dephosphorylation.  For this reason,
many different technologies, including
various colorimetric assays, exist to detect
these enzymes and their activity.
para-Nitrophenylphosphate
Assay Mechanism
para-Nitrophenylphosphate (pNPP) is
widely used as a synthetic substrate to
measure the catalytic activity of various
phosphatases.
The phosphate group of pNPP is cleaved
by the enzyme to yield p-nitrophenol, which
is also colorless since the wavelength
maximum for the electronic excitation of pnitrophenol
in water is 318 nm. However,
under alkaline conditions, the p-nitrophenol
is converted to the p-nitrophenolate anion,
resulting in a bathochromic shift (a change
in absorbance of a compound to a longer
wavelength based on solution conditions)
to around 400 nm. This is at the blue edge
of the electromagnetic spectrum, but since
xxix
we see the color of the reflected light
(as opposed to the absorbed light), the
solution looks yellow.
Things to Keep in Mind…
One of the great selling points of the
pNPP assay is its simplicity, so it’s easy
to see why one might forget that this
bathochromic (a great trivia word!) shift
is a crucial element to the assay.
What if, for example, the enzymatic
activity requires an optimum pH in the
acidic range, as so happens to be the
case with acid phosphatase? 
In that instance, the end point addition of
a strong base (such as sodium hydroxide
or potassium hydroxide) is necessary in
order to obtain the desired yellow color.
Malachite Green
Assay Mechanism
The para-nitrophenylphosphate assay is
great, but what if your favorite
phosphatase doesn’t efficiently
metabolize pNPP? Another commercially
available phosphatase assay is the
malachite green assay. This simple
assay method is based on the complex
formed between malachite green,
ammonium molybdate, and free
orthophosphate (aka inorganic phosphate,
Pi) under acidic conditions.
Orthophosphate, liberated from a
phosphorlyated substrate upon cleavage
by the phosphatase, forms a complex with
ammonium molybdate in a solution of
sulfuric acid.  The formation of the
malachite green phosphomolybdate
complex, measured at 620-650 nm, is
accordingly directly related to the free
orthophosphate concentration. Therefore, it
is possible to quantify phosphorylation and
phosphate release from protein
phosphatase substrates.
Things to Keep in Mind…
This assay measures only inorganic free
phosphate; organophosphates (lipid-bound
or protein-bound phosphates) must first be
hydrolyzed and neutralized prior to
measurement.  Knowledge of the different
classes of organophosphates and their
respective different free energies of
hydrolysis aids in the optimization of assay
conditions.  Higher-energy
organophosphates (acetal phosphates,
xxx
para-Nitrophenylphosphate: From Colorless to Yellow
phosphoanhydrides, etc) possess acidlabile
phosphate groups that can be
released into solution by incubation at
low pH. On the other hand, lowerenergy
organophosphates
(phosphoesters) are stable in acidic
solution and require much harsher
conditions (such as thermal
decomposition) to be detected by this
method.
In the presence of a large excess of
malachite green, the 3:1 ion associate
((MG+)3(PMo12O40 3-)) can easily
form and precipitate in the acidic
aqueous solution. To stabilize the 1:1
ion associate in the aqueous solution,
polyvinylalcohol is added to the solution.
Another thing to take into consideration
during the assay is the possibility of any
redox reactions that could interfere with
the assay. Molybdenum is a transition
metal and so exists in many oxidation
states. In the molybdate anion, it has an
oxidation state of +6.  Reduction of the
acidified Mo(VI) solution by organic
compounds, like ascorbic acid and
reducing sugars (i.e. glucose), or by
inorganic compounds, like SnCl2,
creates the Mo(IV) species, which is blue
in color.
There’s much more where
those came from
These two examples barely skim the
surface of the colorimetric assays. Just
think of all the catalase assays for the
detection of hydrogen peroxide (like
Sigma’s quioneimine derived assay), and
of course, the very popular MTT-based
assays for measuring cytotoxicity.  A better
understanding of the chemistry involved in
these assays helps to provide a deeper
understanding of the reactions, as well as
any required troubleshooting procedures.
Colorimetric assays are just one of the
many ways in which chemistry and biology
work hand-in-hand in scientific research.  I
think it’s fair to say that the multidisciplinary
approach is now recognized as
a vital way to move forward in the drug
discovery process.
My chemical biology professor commented
that in the ‘olden days’ it was enough to
study medicinal chemistry and that the
pharmaceutical companies would teach
you the rest. Now, medicinal chemists are
xxxi
“No sleep ‘til 640nm”, By Malachite Green
expected to understand pharmacology,
cell biology, and molecular biology
techniques.  It takes an entire team of
people, involving many specialities, in
order to be successful in developing
safe and effective drugs.  But it is not
enough simply to involve many
collaborators; I would argue that
researchers on all sides need to learn
to occasionally think alike.  So hopefully
the next time my aforementioned
colleague performs a colorimetric assay
she can don her ‘chemistry hat’ in
addition to her ‘molecular biology hat’,
and remember that drug research is an
interdisciplinary art.
Want More Chemistry for Biologists?
Take a look at this free webinar series from Peter Kirsop at BitesizeBio.com
Chemistry for Biologists
Covered in this series:
Biological Organic Molecules
The Importance of Stereochemistry in Biological Molecules
Features of Organic Molecules
xxxii
How Lab Grade Water is
Purified
“...in practice a number of techniques are used, each of which can
remove a different set of impurities.”
Section 09 By Nick Oswald
There’s something in the water, and it
would love to go after your
experiments.
Straight out of the tap, water contains
microorganisms, endotoxins, DNase
and RNase, salts and other impurities
that could gobble up your experiment in
one bite.
Of course we avoid this drama
completely by using purified water from
which these nasties have been
removed. But how is this purification
done?
Well in practice a number of techniques
are used, each of which can remove a
different set of impurities.
So here are the
techniques:
1. Distillation. A technology as old as
the hills (or at least as the stills that
were hidden in the hills). Water is
heated to its boiling point then
condensed back to liquid. This will
remove many impurities but impurities
with a boiling point equal to or less than
that of water will also be carried over in
to the distillate.
2. Microfiltration. In this technique,
pressure is used to force the water
through a filter with pore sizes of 1 to 0.1
micron in order to remove particulate
matter. Filter diameters lower than 0.2
micron removes bacteria – so-called cold
sterilization.
3. Ultrafiltration uses even smaller pore
sizes (down to 0.003 micron). These are
essentially molecular sieves, which
remove molecules with a diameter larger
than the pore size. It can be used to
remove viruses, endotoxins, RNase and
DNase
4. Reverse osmosis. If you thought that
ultrafiltration used impressively small
pore sizes, you’ll be even more
impressed by reverse osmosis. Reverse
osmosis filters have pore sizes of less
than 0.001 microns, which allows them
to sieve ions depending on their
diameter. This is used for desalting the
water.
5. Filtration through a bed of activated
carbon is useful for removing things like
chloride ions and organic compounds,
which are adsorbed onto the surface of the
carbon.
6. UV radiation. We all know what UV
radiation, at specific wavelengths, can do
to DNA and microorganisms. So UV is an
obvious way to remove microorganisms
from the water. It can also clean up the
water by breaking down certain organic
compounds into less harmful products.
7. Deionization/ Ion exchange. This
technique removes ions from the water by
passing it through a resin bed containing a
mixture of cationic and anionic resins.
Positive ions in the water are attracted to
the anionic resin particles and negative
ions are (yes, you’ve guessed it) attracted
to the cationic resins. The result is that
nicely deionised water comes out of the
other end of the resin bed.
Commercially available water, or water
purification systems will typically use a
combination of these. The higher the water
purity grade, the more techniques used.
xxxiv
(Exactly) Why Enzymes
Have Optimal Temperatures
“Actually it’s reassuringly simple...”
Section 10 By Nick Oswald
Every biologist is familiar with the
profile of the rate of an enzymatic
reaction versus temperature as shown
in the figure.
We know that enzymes from E.coli or
warm-blooded animals tend to have an
optimum around 37°C while those from
thermal vent bacteria have much higher
optimal temperatures.
Surprisingly, I find that many biologists
don’t have a grasp of why enzymes
have these temperature profiles.
Actually it’s reassuringly simple.
Chemists have a rule of thumb that a
10°C increase in temperature gives a
doubling of the reaction rate. This rule
is loosely derived from the Arrhenius
equation.
Basically, as the temperature increases
so does the kinetic energy of the
reactants. This increased kinetic energy
means that the reactants are more
likely to collide with enough energy to
allow the reaction to occur, so the
higher the temperature, the higher the
reaction rate.
Going Up
The first part of the reaction rate profile
(shown below in green), where the rate
is increasing with the temperature,
follows the Arrhenius equation. If the
enzyme was completely stable even at
high temperatures the reaction rate
would continue to increase with
temperature until something else
happened, like one of the reactants
became limiting for example.
Plateauing...
The reaction rate begins to plateau then fall
in the yellow highlighted section of the
graph. This is due to the temperature
approaching the point at which the enzyme
begins to denature (and therefore lose
activity).
xxxvi
A typical plot of enzymes optimal temperature (rate vs temperature). The shape is
caused by competition between the rate increasing with temperature (in line with
Arrhenius equation) vs the breakdown of the enzyme at higher temperatures.
Going down
At even higher temperatures (the red
section) the enzyme is fully denatured
and no activity remains.
The temperature at which the
denaturation occurs is dependent on
the structure of the enzyme, which in
turn is related to it’s evolutionary origin.
Thus, E.coli enzymes have evolved to
cope with temperatures of around 37°C
while enzymes from thermal vent
bacteria have been forced to evolve in
such a way that they can remain stable
at far higher temperatures.
So an enzyme’s optimal temperature is
a trade-off between the Arrhenius-type
dependence on temperature (the hotter
the reaction, the faster the rate) and the
instability of the enzyme as it
approaches then reaches its
denaturation temperature.
Did you find this useful?
Visit BitesizeBio.com for many more
eBooks, articles, webinars and videos on
topics just like this one.
xxxvii
© 2012 Science Squared Ltd, UK
Photos: Bigstockphoto.com, Sxc.hu
xxxviii