Isolation of plasmid DNA
Isolation of DNA/RNA from a bacterial cell extract using phenol extraction
A traditional way of separating the genetic material from a bacterial cell is phenol extraction.
The basic phenol extraction procedure is illustrated in figure 1.
Figure 1. a) The first step of the phenol extraction procedure is to disrupt the cell wall. b) The
cell extract is centrifuged to remove cell debris and only DNA, RNA and protein remains. Phenol
is then added to separate proteins from the genetic material.
As described in figure 1a the first step is disruption of the bacterial cell wall. The solution is then
be centrifuged to remove cell debris. The remaining cell lysate contains DNA, RNA and protein
(1b). When the cell extract is mixed with phenol, the phenol causes the proteins to coagulate.
The aqueous and phenol phases are immiscible and two phases will form. The genetic material,
DNA and RNA, will stay in the aqueous phase and the coagulated protein in the interphase. At a
pH around 7, the genetic material favours the aqueous phase due to the negative charge on the
phosphate groups in the molecules that make them hydrophilic.
1
When the genetic material has been isolated DNA still needs to be separated from the RNA.
This can be done by changing the pH of the solution. Figure 2 illustrates what happens when
the pH is changed.
Figure 2. DNA and RNA can be separated by lowering the pH. DNA will become neutralised and
move to the phenol phase at a pH below 5, due to the fact that the pKa of DNA is higher than
that of RNA. RNA will stay in the aqueous phase.
As described in figure 2, DNA will move into the organic phenol phase at a pH below 5, while
the RNA will remain in the aqueous phase. This is due to a small difference in the sugars in the
DNA and RNA backbones. The sugar in DNA is deoxyribose and in RNA ribose. Ribose has one
extra hydroxyl group compared to deoxyribose, making the RNA molecule more acidic than
DNA. Therefore, the pKa of RNA is lower than that of DNA and DNA will be neutralized first
when the pH is lowered. When DNA is neutralized it becomes hydrophobic and favours the
organic phase, allowing the DNA and RNA to be separated by separating the two phases.
Isolation of DNA/RNA from a bacterial cell extract using chromatographic resins
Isolation of DNA or RNA from a bacterial cell extract using chromatographic resins is a more
modern and convenient way of isolating genetic material. There are many commercially
available kits that can be used for separating plasmid DNA. The ​NucleoSpin​® plasmid isolation
procedure is illustrated in figure 3.
2
The ​NucleoSpin​® kit can be used for small-scale
experiments and high purity plasmids are obtained
rapidly and it is optimized for ​E. coli. The ​NucleoSpin​®
procedure starts with cultivating and harvesting
bacterial cells. The next step is alkaline lysis to remove
genomic DNA and clarification of the lysate. The lysate is
neutralized with a high-salt buffer to create appropriate
conditions for binding of plasmid DNA to the silica
column material. After allowing plasmid DNA to bind to
the column material, the column is washed to remove
any impurities and the last step is elution of the plasmid
DNA using a slightly alkaline buffer. This method is
simple to use and another advantage is that the
obtained plasmids can be directly used for PCR,
transformation, sequencing or any other procedure.
It is important that all genomic DNA is removed during
the purification to obtain pure plasmid DNA. This can be
achieved using alkaline lysis. The principle of the alkaline
lysis is described in more detail in figure 4.
Figure 4. ​The effect of pH on genomic and plasmid DNA and the principle of alkaline lysis.
Alkaline lysis is based on the different properties of genomic and plasmid DNA. The genomic
DNA is linear and longer than the smaller supercoiled plasmids. At a high pH of 12.5, both
3
genomic and plasmid DNA will denature. When the pH is lowered to 7, the plasmid DNA will
re-anneal and go back into its original conformation. This will not happen to genomic DNA as
the linear strands are too long to re-anneal. The DNA will remain a tangled mass that can be
separated from plasmid DNA by centrifugation.
Plasmid DNA purification by anion-exchange chromatography
Purification of DNA can also be achieved through the use
of anion-exchange resins in a chromatographic column.
These positively charged particles (such as DEAE or
diethylaminoethanol) will attach to negatively charged
organic molecules, and detach only when a certain salt
concentration is reached. This principle can be utilized to
achieve
gradual
release
relating
to
growing
electronegativity, allowing for the separation of DNA,
RNA and other organic molecules from each other (Figure
5).
The exact salt concentration at which a specific organic
compound will detach from a resin depends on the pH
inside the chromatography column. pH influences the
hydrogen ion activity, which in turn influences resin
function, as Figure 6 suggests. More H(+) groups will lead
to more positively charged resin molecules, which will
lead to an increased minimum level of salt required for
anion detachment.
Figure 7 illustrates how different organic molecule groups
are obtained at specific NaCl concentrations. Notice how
this graph is specific to pH 7. Should the pH be increased
to 8.5, for example, the Plasmid DNA would be detached
4
at approximately 0.9 M NaCl, which is a considerable difference compared to 1.5 M NaCl.
By using this kind of concentration graph, it is possible to predict the concentration at which for
example specific oligonucleotides or RNAs are eluted.
Figure 7. ​Salt concentrations at which specific organic compounds are released at ph 7.0 by
QIAGEN resins.
Plasmid purification via chromatography matrices
A common way of purifying plasmids is with the help of chromatography matrices, or gel
membranes. Lysate in buffer flows through a silica gel membrane, for example inside a
centrifuge. If pH and salt (eg. CTAB) concentration is optimal, nucleic acids will bind to the
membrane. After a washing step the nucleic acids are detached by using a suitable eluent, and
purified plasmids collected.
5
References
QIAGEN, ​Small-Scale Plasmid Purification — Product Overview, Available:
https://www.qiagen.com/us/resources/technologies/plasmid-resource-center/small-scale%20p
lasmid%20purification%20product%20overview/​ [31.10.2016].
MACHEREY-NAGEL, ​NucleoSpin​®​ Plasmid, Available:
http://www.mn-net.com/tabid/1379/default.aspx​ [31.10.2016].
6
PLASMIDS
Ilona Leppänen
Iiris Hakaste
Plasmid is a molecule of double-stranded DNA that is independent from the cell’s chromosome.
Plasmids are usually circular and can replicate independently. Typically, plasmids can be found
from bacteria and lower eukaryotes. Plasmids are used as integrative and replicative vectors in
molecular biology.
pUC19
The most common cloning vector plasmid used is pUC19. The popularity is based on it’s properties
as a vector molecule. pUC19 is a circular, double-stranded plasmid consisted of 2686 base pairs.
The plasmid contains an ampicillin resistance gene as a selectable marker (picture 1). It also has a
polylinker site which contains restriction sites for many different restriction enzymes. This makes it
possible to cut the plasmid with several different restriction enzymes and thus to have many
different sticky ends or blunt ends to work with.
Picture 1. pUC19 vector plasmid
Maybe the most notable feature of pUC19 is that bacteria colonies grow with different colours
depending on if they have recombinant DNA inserted to the plasmid or not: cells with insert form
white colonies and cells without form blue. This happens because the pUC19 also contains a lacZ
gene, that encodes for part of Beta-galactosidase. The polylinker site is located in the middle of
this gene, as seen in picture 1. When foreign DNA is inserted to the polylinker site, it inactivates
the lacZ gene. The lacZ gene is important factor in the color change, because beta-galactosidase
hydrolyses X-gal, which causes the blue color of the colony. When lacZ gene is inactive, half of the
beta-galactosidase is missing (picture 2) and it cannot hydrolyse X-gal, and thus the colony appears
with white color. Also IPTG is needed on the growth plate, because it induces lacZ transcription.
Picture 2. The principle of the lacZ complementation
Prokaryotic expression systems
Prokaryotic plasmids usually contain the same basic features. There is a sequence called
prokaryotic promoter (drawn with pink in picture 3), which is the starting spot for transcription.
After that there is a sequence that works as ribosome binding site. Then follows the polylinker or
multiple cloning site (MCS), that contains from a few to many different restriction sites for
restriction enzymes. Then we have the transcription termination site. Plasmids also contain the
sequence called origin of replication: this is a prerequisite for the plasmid to be able to replicate
independently from the cell’s chromosome. The plasmids usually contain also the prokaryotic
selectable marker, which in this case is the ampicillin resistance gene.
Picture 3. Prokaryotic plasmid
Eukaryotic Expression vectors
An eukaryotic expression vector has the same features as any vector has. These include origin of
replication, selectable marker and a multiple cloning site. The origin of replication is the gene
sequence where the replication is initiated. The selectable marker is often an antibiotic resistance
gene. These genes are a type of reporter gene that indicate whether the host has taken up and
expressed the genetic material. The multiple cloning site is the transcription unit, which includes
the promoter, MCS for the gene of interest and a DNA segment with both termination and
polyadenylation signals. Polyadenylation means addition of poly(A) tail to a mRNA to produce
mature mRNA for translation. The multiple cloning site, also called as a polylinker, contains
multiple restriction sites. Restriction enzymes recognize these sites and cut the plasmid from there
so that the plasmid can be inserted into the host DNA.
The eukaryotic expression vector contains origin of replication for both E. coli and
eukaryotic cell and additionally a selectable marker for the eukaryotic cell (ESM) and selectable
marker for E. coli the Ampr. The selectable marker and ori for E. coli are incorporated in the vector
so that they can be manipulated and amplified in the bacterium. These processes are easier to do
in a less complex environment.
Picture 4. Eukaryotic expression vector
Yeast artificial chromosome
Yeast artificial chromosomes also known as YACs are genetically engineered chromosomes derived
from the DNA of the yeast, Saccharomyces cerevisiae, which is then ligated into a bacterial
plasmid. The YAC is designed to clone large fragments of DNA. Up to 1 million base pairs can be
inserted into YACs. YACs are plasmid shuttle vectors, which means it can propagate in two
different host species. These vectors are capable of replicating and being selected in hosts such as
E. coli. The YACs are relatively small and circular when amplified but become linear and large when
they are introduced as cloning vector in yeasts.
The construction of a YAc is fairly complex as it has various different features. It has
both vector and chromosome features. The YAC contains restriction sites where two restriction
enzymes (SmaI and BamHI) cleave the plasmid and break it up into two DNA arms. These arms are
ligated with the DNA insert (>100kb). The ARS sequence is an autonomously replicating sequence,
which is necessary for replication and provides the origin of replication. The centromere (CEN),
which is a part of the chromosome, is for segregation at cell division. The two telomere sequences
(TEL) at the end of the linear YAC are required to insure stability of YAC ends. The URA3 and TRP1
are selectable markers, one for each arm.
The yeast cells take up the YAC and the inserted DNA. The YAC is maintained as a
separate chromosome in the host cell and is highly stable. When the yeast cell grows and divides
they amplify the YAC DNA, which can be then isolated and used for various applications. These
applications include for example the physical mapping of the human genome, analysis of large
transcription units and formation of genomic libraries containing DNA from individual human
chromosomes.
Picture 5. Yeast Artificial Chromosome
Integrative plasmids
In order for a plasmid to replicate independently within a cell it must posses a sequence of DNA
that acts as the origin of replication. Some plasmids lack the ori and can be incorporated into the
host chromosome. They integrate with the host chromosome. Integrative plasmids are referred to
episomes in prokaryotic cells. Also yeast-integrating plasmids (Yip) lack an ori and must be
integrated into the host chromosome via homologous recombination. What was clearly stated in
the lecture was that the integrative plasmid may be replicated and stably maintained in the cell
through multiple generations and in cannot in any stage exist as an independent plasmid.
CHEM-E81120 Laboratory Course in
Biosystems and Biomaterials Engineering
Principles of Polymer Chain Reaction
and in vitro DNA Amplification
Maisa Vuorte & Henriikka Vekuri
30.10.2016
Contents
1 Introduction
1
2 Basic principles of PCR
1
3 Primer Design
4
4 DNA Polymerases
5
5 Applications of PCR
7
1
Introduction
The polymerase chain reaction (PCR) was developed by Kary Mullis in the
1980s for amplification of DNA. 1 PCR enables rapid and specific in vitro
replication of DNA sequences typically ranging from 0.1 to 10 kbp. Starting
from possibly a whole genome, selected regions of the DNA or RNA template
can be replicated exponentially in relatively short time window. 2;3 PCR has
a constantly expanding, wide number of applications in many fields including
the biotechnological and pharmaceutical industry, medicine, forensic sciences
and sequencing technologies 2;4;5 . The process enables rapid amplification of
DNA samples for example sequencing, DNA fingerprinting, and diagnosis of
hereditary diseases.
In this report, we aim to cover the basic principles of the polymerase
chain reaction as well as possible manipulation of the reaction environment
for introduction of purposeful mutations. In addition to covering the biochemical principles of PCR, we introduce the basic rules of quality primer
design. PCR aims for specific and high-yield amplification of the target DNA
sequence, which in turn requires good primers. We also provide a breakdown
of the general biological background and function of commonly used DNA
polymerases in PCR. Finally, we summarize the most common applications
of PCR including DNA fingerprinting, phylogenetics, as well as the introduction of mutations or deletions through PCR.
2
Basic principles of PCR
PCR or the polymerase chain reaction is used for exponential amplification of
a small number of target DNA molecules. Mainly due to its high specificity
and sensitivity, the method is preferred for replicating small sample sizes of
target DNA for use in a wide variety of applications. PCR relies mainly on
thermal cycling of the reaction mix consisting of a small amount of sample
DNA, a mixture of the four nucleotides A,T,C, and G, a pair of primers, and a
thermostable DNA polymerase suspended in a buffer containing monovalent
potassium ions and bivalent cations, such as magnesium or manganese. 1;2
The sample DNA contains the target region that is amplified through
PCR. This is also called the template. Through incorporation of a reverse
transcriptase active enzyme, an RNA template can be used in place of a
typical DNA template. In this case, the RNA template is first transcribed
into DNA prior to PCR amplification. An example of a reverse transciptase
active DNA polymerase is the Tth DNA Polymerase, whose reverse transcriptase activity can be controlled through the presence of Mn2+ in the reaction
1
buffer.
Primers are short, single stranded DNA sequences that bind to the DNA
template and act as a starting point for strand replication by the DNA polymerase. The two primers are hybridize to the beginning and end on the
target region of the DNA template and are complementary to the sense and
anti-sense strands respectively 6;2 . Thus the primers drive DNA replication
towards each other. This set up is illustrated in figure 1(a). Principles of
good primer design are covered in chapter 3 of this report. The complementary strands are synthesized by the DNA polymerase. Due to the elevated
temperatures used during a typical PCR cycle, a thermostable variant is preferred 1 . The characteristics of DNA polymerases used in PCR are discussed
in more detail in chapter 4 of this report.
The role of deoxynucleoside triphosphates (dNTPs) is to act as building
blocks for the synthesized DNA strands. Typically, an equimolar mixture of
each of the four dNTPs is used for PCR. However, certain mutations in the
PCR product can be favored by unbalancing the dNTP mixture.
The buffer solution used in PCR contains ions that are essential for a
successful PCR reaction. Positively charged potassium ions are added to the
buffec in the for of KCl. The K+ ions work to effectively screen the negative charges in the DNA backbone, thus providing increased stability for the
primer-template complex. Increased KCl concentration has been shown to
slow down the denaturation of long DNA strands. 7 Bivalent cations, such
as manganese, can affect the efficiency and sensitivity of the PCR reaction.
DNA polymerases tend to require bivalent cations as co-factors for activity.
dNTPs and oligonucleotides also bind to Mn2+ ions in equimolar concentration. As such the concentration of bivalent cations must exceed that of
phosphate groups contributed by both the dNTPs and oligonucleotides. It
has been found that excessively high concentrations Mn2+ ions in the buffer
lower the fidelity of DNA polymerases such as Taq polymerase. This effect
is utilized in error prone PCR 8 .
PCR relies on repeated thermal cycling of the reaction mix. A typical
PCR cycle, as illustrated in figure 1(a), consists of three distinct phases: denaturation, annealing of primers, and extension of complementary strands 2 .
During the first phase of the cycle, the reaction mix is heated to approximately 90 ◦ C. The increased temperature causes the hydrogen bonds between
the complementary strands of the double helix to break. This action causes
the double stranded structure of the sample DNA to denaturate. After denaturation, the temperature is lowered to 50 − 65 ◦ C. At this temperature
the primer sequences anneal to the template DNA. The temperature during
the annealing phase of the PCR cycle must be low enough to accommodate hybridization and hydrogen bond formation but high enough to allow
2
(a)
(b)
Figure 1: a) The PCR cycle includes three distinct steps carried out at different
temperatures. First the reaction mixture is heated to ≈ 90 ◦ C to denaturate the
double stranded structure of the DNA. After denaturation, temperature is lowered
to 50 − 65 ◦ C for specific hybridization of the primers to the target DNA sequence. The temperature is again raised to the optimum temperature of the DNA
polymerase used in the reaction (≈ 70 ◦ C) and the complementary strand to the
target DNA is synthesized. b) PCR enables specific and rapid amplification of target DNA sequence. The amount of target DNA doubles after each cycle, resulting
in exponential amplification.
only specific annealing, leaving no room for mismatches between base pairs.
After annealing of the primers, the temperature is raised to the optimum
3
temperature of the DNA polymerase, typically at around 70 ◦ C. During
the elongation phase, complementary strands are synthesized in the 5’ → 3’
direction. During the next PCR cycle, the freshly synthesized strands act
as templates. This leads to doubling of the amount of template DNA after
each consecutive cycle. This process is know as exponential amplification. 2
It is important to check that correct regions of the template were amplified
during PCR. This can be done by separating the different PCR products by
size using gel electrophoresis.
3
Primer Design
Specific and high-yield PCR amplification of target DNA or RNA requires
good primers and well planned primer design is essential for a successful
reaction. Primers are short single stranded DNA sequences that bind to the
target region of the sample DNA during the second step of the PCR cycle,
which is illustrated in figure 1(a). This step is often called the annealing
step of the PCR cycle 2 . A PCR reaction requires two primers: a forward
primer that hybridizes to the 3’-end of sense strand of the template gene
and a reverse complement primer that hybridizes to the the 3’-end of the
antisense strand of the template gene. This set up is illustrated in figure 1(a).
In addition to ensuring specific amplification of the desired PCR product,
enzymatic restriction sites can be designed as a part of the primers and thus
incorporated into the product. 6;9
The length of a primer affects its binding specificity as well as its binding
speed. The primer must be long enough to assure specific binding as well as
short enough for easy and speedy hybridization to the template. The length
of a primer typically varies from 18 to 30 bp 6 .In addition to the length
of the primers, specific hybridization is controlled by the temperature of the
annealing step of the PCR cycle as has been formerly described in this report.
Specific and strong binding of the primers to the template DNA is encouraged by the high GC-content of the primers, typically ranging from 40
to 60%. The GC content of the primer determines the primer melting temperature Tm . Tm is characterized as the temperature at which one half of the
DNA duplex will dissociate to become single stranded. This is used as an indicator of duplex stability. Commonly primers with Tm at around 52− 58 ◦ C
are preferred. The Tm of a primer is calculated according to the following
formula 9 :
Tm = 2 ◦ C × (NA + NT ) + 4 ◦ C × (NC + NG )
(1)
where NA , NT , NC , and NG denote the number of adenine, thymine, cytosine,
and guanine in the primer respectively. The Tm of the chosen primers for the
4
PCR reaction determines the annealing temperature Ta of the PCR cycle.
Typically the annealing temperature chosen is around 5 ◦ C bellow the Tm .
The thermostable DNA polymerase used in the PCR reaction catalyzes
the extension of the 3’ end of the complementary DNA strand. As such, the 3’
end can be considered the most crucial part of the primer. It is recommended
to have G and C bases within the last five bases of the 3’-end of the primer
to ensure specific hybridization. This is known as the GC clamp. However,
more than three G or C bases at the 3’-end should be avoided 9 .
Primers must also be screened for possible unwanted secondary structures, such as hairpins, self-dimers and cross-dimers 6 . Hairpin structures are
formed through intramolecular interaction of the primer while self-dimers
and cross-dimers are formed intermolecularly by homologous primers. To
avoid the formation of undesired secondary structures, homology within and
between the primers, as well as multiple, consecutive repeats of a single nucleotide or a dinucleotide should be avoided when designing primers. To
avoid unwanted PCR products, it is also important that the chosen primers
are not compatible with any other regions in the sample genome. This crosshomology can be investigated through, for example, BLAST 10 .
4
DNA Polymerases
Because of the high temperatures used in the PCR reaction, a thermostable
DNA polymerase is required. Proteins tend to denature at high temperatures, resulting in loss of function in enzymes. Previously, the so called
Klenow fragment isolated from Escherichia coli was the default DNA polymerase in PCR reactions. The Klenow fragment exhibits 5’ → 3’ polymerase
activity and 3’ → 5’ exonuclease activity. However, due to its thermosensitive
nature, more enzyme is required to be added after each PCR cycle. Modern
thermostable variants of DNA polymerase have been originally isolated from
thermophilic and hyperthermophilic bacteria and archaea. Typically polymerases with an optimum temperature at around 70 ◦ C are preferred. DNA
polymerases isolated from different organisms have different kind of properties regarding molecular weight, half-life, proof reading, error rate, extension
rate, reverse transcriptase activity and polyadenylation. The proper polymerase can be decided for each experiment after considering the necessary
and unnecessary properties of the polymerase for each specific experiment.
Taq DNA polymerase is isolated from the thermophilic bacterium Thermus aquaticus, as illustrated in figure 2. It does not have 3’ → 5’ exonuclease
activity which is the ability of proof reading and this leads to an error rate
of 2.28 × 10−5 . At 95 ◦ C it is thermostable for 40 minutes. On the other
5
Figure 2: Thermus aquaticus bacteria deposited on a Millipore filter of 0.22 µm.
PCR utilizes thermostable DNA polymerase, often with optimum temperature at
around 70 ◦ C. Taq polymerase was originally isolated from the thermophilic bacterium Thermus aquaticus.
hand, it is relatively fast and performs polyadenylation on the 3’ ends which
results in 3’ A overhangs which can be useful for some experiments. Phusion
is a pyrococcus-like enzyme with a processivity enhancing domain. It does
proofreading; it recognizes incorrectly matched base pairs, returns one base
pair backwards and corrects the mismatch. Because of this, its error rate
is much lower than that of Taq: 4.4 × 10−7 . Its half-life at 95 ◦ C is over 6
hours. Phusion does not add adenines at the 3’ end which results in blunt
DNA ends. Compared to Taq it is slower. 11 Properties of phusion and taq
are gathered and compared in figure 3.
Figure 3: Phusion, a pyrococcus-like enzyme with a processivity enhancing domain,
compared to Taq, a DNA polymerase isolated from Thermus aquaticus.
6
5
Applications of PCR
PCR has applications in many fields including medicine, forensics and research. Medical applications include for example genetic testing. DNA samples can be analyzed for disease causing mutations and prenatal testing. Also
when using in vitro fertilization cells can be tested for mutations before implantation. PCR can aid tissue typing for organ transplantation instead of
using tests based on antibodies. Oncogenes can be identified which can help
providing individualized therapy for cancer patients. PCR is also needed for
diagnostic microbiology. Culturing, microscopy and other visual detection
methods can be troublesome and also yield unreliable results. Some microorganisms cannot even be cultured or they grow very slowly. Diagnostics
based on DNA sequences, when performed correctly, are highly specific. The
key is to select the primers specific for the genome or gene of interest. 12
Every individual (except for identical twins) has a unique genome. This
is the basis for DNA fingerprinting. PCR can be applied in forensics in
differentiating individuals. Even a small sample from a crime scene can be
enough as long as it contains enough DNA, theoretically one strand, for PCR
amplification. It can be compared to suspects and used for identifying the
correct one. A similar method can be applied for paternity testing. 13
Several research fields rely on molecular genetics which usually requires
PCR. PCR is handy when DNA is needed for sequencing purposes. Only
the neighbouring sequences of the gene or genes of interest have to be known
for designing the primers. Phylogenetic studies study evolutionary history
and relationships between different organisms and species. PCR can be used
to amplify 16S rRNA genes that contain sequences well conserved throughout evolution that can be used for targeting primers. 16S rRNA genes also
contain hypervariable sequences that are species specific and they can be
used for differentiating between species. Also small samples from ancient or
fossilized plants or animals can be amplified and cloned with PCR. 12;14
Different PCR applications come back to the design of primers. For example, two separate PCR reactions can be performed with primers that have
overhangs complementary to the end of the DNA molecule in the other PCR
reaction. Thus, the two DNA molecules are extended with a new sequence.
Primers are removed and the two amplified DNA molecules are mixed together with new primers targeting the far ends. When a third PCR is performed the two sequences are fused. Overlapping PCR can also be used to
create deletions.
Directed PCR mutagenesis targeting a plasmid utilizes primers that include wanted mutations. First, linear PCR is performed with Pfu polymerase. Secondly, the templates are digested with the restriction enzyme
7
Dpnl that recognizes methylated sites. Thus, the templates are digested but
the newly synthesized plasmids containing mutations remain intact. Because
DNA polymerase cannot complete the full plasmid, nicks are left in the plasmid. When the plasmids are transformed the nicks are repaired in vivo by a
ligase. 15
8
References
[1] R. K. Saiki, D. H. Gelfand, S. Stoffel, S. J. Scharf, R. Higuchi, G. T.
Horn, K. B. Mullis, and H. A. Erlich. Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science,
239(4839):487–491, Jan 1988.
[2] B. Alberts. Molecular biology of the cell. Garland Science, 2008.
[3] D. C. Rio. Reverse transcription-polymerase chain reaction. Cold Spring
Harb Protoc, 2014(11):1207–1216, Nov 2014.
[4] G. Muyzer, E. C. de Waal, and A. G. Uitterlinden. Profiling of complex
microbial populations by denaturing gradient gel electrophoresis analysis
of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl.
Environ. Microbiol., 59(3):695–700, Mar 1993.
[5] J. L. Weber and P. E. May. Abundant class of human DNA polymorphisms which can be typed using the polymerase chain reaction. Am.
J. Hum. Genet., 44(3):388–396, Mar 1989.
[6] PREMIER Biosoft. PCR primer design guidelines.
[7] S. Cheng, Y. Chen, J. A. Monforte, R. Higuchi, and B. Van Houten.
Template integrity is essential for PCR amplification of 20- to 30-kb
sequences from genomic DNA. PCR Methods Appl., 4(5):294–298, Apr
1995.
[8] D. S. Wilson and A. D. Keefe. Random mutagenesis by PCR. Curr
Protoc Mol Biol, Chapter 8:Unit8.3, May 2001.
[9] European Molecular Biology Laboratory. Cloning: PCR strategy, primer
design.
[10] S. F. Altschul, W. Gish, W. Miller, E. W. Myers, and D. J. Lipman.
Basic local alignment search tool. J. Mol. Biol., 215(3):403–410, Oct
1990.
[11] New England Biolabs. Thermophilic dna polymerases.
[12] M. A. Valones, R. L. Guimaraes, L. A. Brandao, P. R. de Souza,
A. de Albuquerque Tavares Carvalho, and S. Crovela. Principles and
applications of polymerase chain reaction in medical diagnostic fields: a
review. Braz. J. Microbiol., 40(1):1–11, Jan 2009.
9
[13] R. Decorte and J. J. Cassiman. Forensic medicine and the polymerase
chain reaction technique. J. Med. Genet., 30(8):625–633, Aug 1993.
[14] A. Klindworth, E. Pruesse, T. Schweer, J. Peplies, C. Quast, M. Horn,
and F. O. Glockner. Evaluation of general 16S ribosomal RNA gene
PCR primers for classical and next-generation sequencing-based diversity studies. Nucleic Acids Res., 41(1):e1, Jan 2013.
[15] A. V. Bryksin and I. Matsumura. Overlap extension PCR cloning: a
simple and reliable way to create recombinant plasmids. BioTechniques,
48(6):463–465, Jun 2010.
10
CHEM-E8110 - Laboratory Course in Biosystems and Biomaterials Engineering
Keivan Khademi Kalantari – [email protected]
Nguyen Hoang Vu - [email protected]
Assignment
RESTRICTION ENZYMES
I.
Overview: Biological origin and function of restriction enzymes
Restriction enzymes, first and foremost, are the tools bacteria came up with to defense
itself from foreign DNA invasion. Restriction enzymes have a recognition domain which
enable them to bind a specific site in the DNA and exert their cleavage function either there or
a random site far away. The final objective is to cut the guest’s DNA into small, harmless
fragments, thereby maintaining host’s genome integrity. The host’s genome is protected from
its own restriction enzymes by means of DNA
methylation, which is catalyzed by DNA
methylases. Restriction enzyme and its
corresponding methylase are referred to as a
restriction-modification system.
With the ability to recognize and cleave at a
specific sequence, restriction enzymes have
endowed biochemists with unprecedented power
for DNA modification. Nowadays, restriction
enzymes are used extensively in: traditional
cloning, DNA mapping, methylation detection, in
vitro DNA assembly, construction of DNA
libraries, creation of nicks in DNA, in vivo gene
editing.
II.
Classification, Mechanism, Structure
Restriction enzymes are categorized according to their
cleavage sites, recognition site, structure, and cofactor
requirement. There are 3 main classes: type I, II and III. Of
these, type II restriction enzymes are most useful in DNA
specific manipulation because they cleave DNA within their
recognition site.
Structure: Most restriction enzymes are homodimers,
with each subunit recognizes and cleaves a single strand of the
double-stranded DNA. Therefore, most recognition sequences
are palindromic.
Recognition sites of restriction enzymes typically have 4-12 nucleotides. The length of
recognition sites has an implication on the size of fragments produced because the longer the
recognition sequence, the more specific the enzymes, as they would cleave at fewer sites.
III.
Isoschizomers
When two or more restriction enzymes have the same recognition sequence and cut in a same
way, they are isoschizomers, e.g SphI (CGTAC/G) and BbuI (CGTAC/G) are isoschizomers,
whereas those restriction enzymes that have a same recognition sequence but cut it in a different
location are called neoschizomers e.g. For example, smal (CCC/GGG) and XmaI (C/CCGGG)
are
neoschizomers
of
each
other.
Despite sharing the same DNA recognition site, isoschizomers may have different optimal
reaction temperatures. Anza 13 EspI has an optimal reaction temperature of 37°C, while its
isoschizomer BsmBI works best at 55°C and has only 20% relative activity at 37°C.
IV.
Blunt / sticky ends
Sticky ends are usually a result when restriction enzyme type II cleaves the DNA in a way
that it leaves 2 short single strand, usually 4-12 nucleotide long, overhangs that are
complementary to each other. This cleavage site makes it possible for complementary single
stranded DNA or RNA to attach easily with weak hydrogen bonds but is later tightened by
ligase forming covalent bonds. This is important method in molecular cloning.
Example:
Blunt ends are formed when other restriction enzymes cleave the DNA at the same location
on both strands of DNA, which results two double stranded DNA without overhangs.
Example:
These two cleaving methods have their own advantages and disadvantages. Sticky end is
useful when we want to easily combine to single stranded DNA, but it is limited with only
complementary strings of DNA. Blunt ends are hard to connect because of the lack of easily
connecting weak hydrogen bonds. Even though ligase enzyme can connect the blunt ends
also, although it is much easier for sticky ends, since they can form a preliminary bond. That
said, the variety of different combination are possible with blunt ends, since it is not limited
by the single stranded DNA that requires a complementary string to form the connection.
V.
Compatible ends
Compatible Cohesive Ends are produced when you have two separate enzymes that
recognize very similar sequences, and cut so that the overhang produced by one can
hydrogen-bond with the overhang produced by the other. The below enzymes show a
case where this is true.
We can also have compatible ends that can’t be cleaved anymore when there is just right
amount of differences between two DNA sequences.
VI.
Star activity
Under non-standard reaction conditions, some restriction enzymes are capable of cleaving
sequences which are similar, but not identical, to their defined recognition sequence. This
altered specificity has been termed “star activity". It has been suggested that star activity is a
general property of restriction endonucleases and that any restriction endonuclease will cleave
no canonical sites under certain extreme conditions.
The manner in which an enzyme's specificity is altered depends on the enzyme and the reaction
conditions which induce star activity. The most common types of altered activity are single
base substitutions, truncation of the outer bases in the recognition sequence, and single-strand
nicking. Some enzymes exhibit relaxation of sequence specificity under standard conditions
and in the presence of the cognate site are capable of cleaving non-cognate (secondary) sites.
Factors that provoke star activity are: >5% glycerol, low ionic strength (NaCl), high pH , high
enzyme: DNA ratio, and using Mn2+ as a cofactor instead of Mg2+.
VII.
Engineering of restriction enzymes
- Engineering improved performance: as mentioned above, some restriction enzymes
exhibit cleavage activity at non-cognate sites which reduce their specificity and increase
unexpected products in bio applications. Naturally occurring enzymes mostly work under a
strict set of conditions which makes them ill-suited for the varying experimental conditions in
laboratory settings. Many research has been directed to solve this problem and give rise to High
Fidelity Restriction Enzymes, which could maintain high specificity under wide range of
reaction conditions.
- New specificities: By altering the recognition site of the Type II restriction enzyme
named MmeI, scientists have been able to engineer a host of new restriction enzymes with new
sequence specificities. An additional advantage of research in this direction is that any
alteration in sequence recognition will not only affect specificity of restriction domain but also
methylation domain.
- Nicking endonucleases: aside from homodimeric enzymes with each monomer cleave
half of a palindromic site, there are also other type of restriction enzymes with have strandspecific nicking property.
VIII.
Application
As outlined above, restriction enzymes have applications in a wide range of fields. Within
the scope of the lab course, we will encounter restriction enzymes in traditional cloning and
synthetic biology
a.
Traditional cloning
In conjunction with DNA ligases, restriction enzymes are the basic tools for the ‘cut and
paste’ procedure in gene cloning to move a DNA fragment from one organism to another.
b.
Synthetic biology is another domain where restriction enzymes have been
extensively used. Three mainstream technologies to create a new artificial biological system
are: BioBrick, Golden Gate Assembly, Gibson Assembly. Of these, BioBrick, which is used in
the lab course, based on the idea of quickly assemble a new biological ‘building’ using
standardized genetic ‘bricks’. Restriction enzymes and ligases are used to glue every piece of
DNA together.
Fall Ligation and Transformation Maija Leppä & Sofia Julin Advances in DNA manipulation approaches over the last decades have opened many new avenues for recombinant DNA technologies in both research and industrial biotechnology. The discovery of DNA ligases and the study of prokaryotic gene transfer has enabled the introduction of exogenous genetic material into cells leading to the creation of recombinant genes and organisms. Therefore, methods such as ligation, transformation and transduction are of high importance in genetic engineering. [ T y p e t h e c o m p a n y a d d r e s s ] 16 Ligation Ligation is the process in which DNA sequences are joined together with covalent phosphodiester
bonds. The enzyme DNA ligase catalyzes the formation of the phosphodiester bond between the 3’
hydroxyl end of one DNA fragment (acceptor) and the 5’phosphate end of another fragment (donor).
The DNA ligation mechanism progresses in three steps and requires energy in the form of either
adenosine triphosphate (ATP) or nicotinamide adenine dinucleotide (NAD+). As seen in Figure 1 the
first step is an adenylylation of the DNA ligase in which the ligase itself reacts with ATP (or NAD+) to
form enzyme-AMP and pyrophosphate (PPi) (or nicotinamide mononucleotide, NMN, in the case of
NAD+). In the second step the AMP is transferred from the DNA ligase to the 5’ phosphate in the nick
of the donor and a pyrophosphate bond is formed. In the final step the 3’ hydroxyl group of the acceptor
attacks the 5’ phosphate, a phosphodiester bond is formed, and the AMP-molecule released. [1][2][3,
pp. 989-990]
Figure 1. Mechanism of the DNA ligase reaction. The DNA ligation is a three-step reaction that needs energy in the form of either ATP
or NAD+. The first step is an adenylylation of the DNA ligase and results in an enzyme-AMP-complex. The AMP is then in the second
step transferred to the 5’ phosphate in the nick and a pyrophosphate bond is formed. In the third step the 3’-hydroxyl group attack the
phosphate, which result in a formation of a phosphodiester bond and a release of the AMP. [3, p. 990]
As already mentioned, either ATP or NAD+ is an absolute cofactor requirement for the ligation
mechanism. ATP is a nucleoside triphosphate composed of an adenine ring, a ribosome sugar and three
phosphate groups. NAD+ on the other hand is a dinucleotide consisting of two nucleotides, one with an
amine base and the other with a nicotinamide base, attached to each other with a phosphoanhydride
bond. Both also consist of an adenosine monophosphate (AMP) that can be used to form the enzyme-
1
AMP-complex in the first step of the DNA ligase reaction. [14, pp. 66-67,72] Also the 5’ phosphate of
the donor is of importance for the ligation mechanism and its removal would inhibit ligation because the
DNA ligase mechanism cannot continue in the second step without it [11].
DNA fragments with sticky ends can with DNA ligase be joined to other sticky end DNA sequences of
different origin if they both have been cleaved with the same restriction enzyme and therefore have
compatible ends. These ligations can also be recut again at the same restriction site if the same
restriction enzyme that initially cleaved the DNA fragments is used. The ligation namely reconstructions
and seals the nick. [4, pp. 196-197]
Ligases There are several commercial ligases available and which DNA ligase to choose depends on the DNA
fragments that are ligated and the cofactors that are available. E. coli DNA ligase, T4 DNA ligase and
T4 RNA ligase are commonly used ligases in recombinant DNA technology. The T4 DNA and RNA
ligases are isolated from E. coli infected with the bacteriophage T4 whereas the E. coli DNA ligase, as
the name implies, is isolated directly from the bacteria Escherichia coli [1, p. 44][2].
Both E. coli and T4 DNA ligases can be used to ligate cohesive, complementary overhangs or nicks in
double stranded DNA, but of these two T4 DNA ligase is the only suitable option for ligation of blunt
ends in double stranded DNA. Another difference between E. coli and T4 DNA ligases is the energy
source they require for the DNA ligation reaction. In contrast to other DNA ligases E.coli DNA ligase
requires NAD+ as a cofactor instead of ATP. [1, pp. 44-47][12] [13] For ligation of single-stranded
RNA and DNA the T4 RNA ligase can be used in the presence of ATP [5].
Dephosphorylation of DNA using phosphates Restriction enzymes cut DNA at specific restriction sites. When these restriction enzymes are used in
plasmids the circular forms are momentarily lost due to the cutting. This leads to a linear plasmid
formation with a free hydroxyl group at the 3'-end and a free phosphate group at the 5’-end. As
discussed earlier the ligation of DNA strands happens when a phosphate group and a hydroxyl group
combine with the help of a ligase and required cofactors such as ATP. [6][2][3]
When inserting DNA fragments into plasmids the goal is to ligate the fragment into cut restriction sites.
However, if a linearized plasmid has a free hydroxyl group at one end and a free phosphate group at the
other end, it can self-circularize without the insert fragment being ligated into the plasmid. To avoid
self-circularization alkaline phosphatases are used. They are hydrolase enzymes that remove phosphate
groups from various molecules. This way the phosphate group at the 5’-end of the linearized plasmid is
replaced with a hydroxyl group. [7]
2
Without a phosphate group at the 5’-end, the plasmid cannot self-circularize. The insert DNA fragment
does, however, have a hydroxyl group at the 3'-end and a phosphate group at the 5’-end, and can
therefore be introduced to the linearized plasmid. A ligase can now from two phosphodiester bonds
between the insert DNA fragment and the ends of the linearized plasmid, forming a circular plasmid.
[7][8]
Even though the addition of phosphatases should ideally remove the option for self-circularization, it
still occurs quite commonly even when phosphatases are present. The use of several restriction enzymes
that cleave the plasmid at several sites, and therefore increase the amount of cuts, makes it, however,
possible that insert DNA fragment is attached to the plasmid at high rate. Also, the removal of ‘stuffer’
fragments increases the attachment of insert DNA. ‘Stuffer’ fragment is a small fragment of DNA that is
replaced by insert DNA fragment. By removing this small fragment it cannot rejoin the passenger DNA
during ligation and decrease the amount of conversion. Figure 2 shows what maximizes the formation of
recombinant molecules. [8]
Figure 2. Maximizing formation of recombinant molecules. The graph shows the correlation between the concentration of termini of
foreign DNA and fraction of vector DNA converted to monomeric plasmid/ foreign DNA chimeras. When a vector is cleaved with more
than one enzyme the conversion is higher than with one enzyme. The removal of ‘stuffer’ fragment increases the conversion as well.
Ligase independent ligation Although ligases are highly used when creating recombinant DNA molecules, their use is not the only
method for attaching DNA fragments. There are several ligase-independent methods, of which TOPO
Cloning is one example. It is a method that utilizes the functions of topoisomerases to attach bluntended PCR products into vectors. The biological role of topoisomerases is to cleave and rejoin supercoiled DNA ends in the replication process. It functions both as a restriction enzyme and as a ligase.
This principle is used to attach DNA fragments to a vector. [9]
The topoisomerase that is used in TOPO Cloning was originally derived from Vaccinia virus. It has a
topoisomerase I that specifically recognizes the pentameric sequence 5´-(C/T)CCTT-3´ and forms a
covalent bond with the phosphate group attached to the 3´ thymidine deoxynucleoside. The
3
topoisomerase cleaves one DNA strand, and the DNA unwinds. The enzyme then ligates the ends of the
cleaved strand back together and releases itself from the DNA. In TOPO Cloning the TOPO vectors are
readily linearized and have the topoisomerase I covalently attached to each 3’ overhanging thymides.
This enables the vectors to ligate DNA sequences with compatible ends. Figure 3 shows a schematic
presentation of the principle of TOPO Cloning. [9]
Figure 3. Principle of TOPO Cloning. Topoisomerase I is attached to the thymides at its recognition sites in the TOPO vector. The
topoisomerase cleaves one strand of the DNA of the PCR product and the four-nucleotide overhang in the vector invades the double-strand
DNA of the PCR product. The PCR product is ligated to the TOPO vector by the topoisomerases that are then released.
Introduction of DNA into cells There are several ways how DNA can be introduced into cells. These are transformation, conjugation
and transduction. These methods are natural horizontal gene transfer among prokaryotes that have been
utilized in genetic engineering. [10]
Transformation is the intake of exogenous genetic material from the environment. The cell must be in a
state of competence, meaning that the cell membrane has gone through changes, and allows the
extracellular DNA to enter the cell. This competence can be acquired either naturally or artificially. In
genetic engineering electroporation is an example of artificially acquiring competence. It is often used to
get recombinant DNA into cells by exposing the target cell to high-voltage electrical pulses, which
makes the cell membrane permeable. Electroporation is fast and cheap but may sometimes be too
damaging for the cells. [10][11]
Conjugation is a horizontal gene transfer not that much utilized in gene engineering but common among
bacteria. In conjugation bacterium with a F (fertility) plasmid forms a conjugation pilus that attaches it
to a bacterium with no F plasmid (F-). A single-stranded DNA of the F plasmid is transferred along the
pilus to the F- cell. When a complementary strand is synthesized the originally F- bacterium becomes
F+ as it now has a F plasmid. In Hfr cells the F-plasmid can integrate itself into the chromosome of the
bacteria. When conjugation pilus is formed the bacterium receiving genetic material receives both
chromosomal DNA as well as plasmid genes. [10][11]
Transduction is a way to introduce DNA from viruses into bacteria. Phages are bacterial viruses that
inject their genome into the host bacteria for replication. In genetic engineering this can be utilized by
4
packing recombinant DNA into a phage to deliver it to the bacteria. The same principle can be utilized
using not only bacterial viruses. In viral transformation recombinant DNA is packaged into a viral
particle and virus delivers DNA into target cell that can be plants or mammalian cells. [10][11]
References [1] Primrose, S. B. & Twyman, R. M. Principles of Gene Manipulation and Genomics. 7th Edition.
Malden: Blackwell Publishing. 2006. 644 p.
[2] Lehman, I.R. DNA ligase: structure, mechanism, and function. Science. Vol. 186:4116. 1974. pp.
790-797. DOI: 10.1126/science.186.4166.790.
[3] Nelson, D. L. & Cox, M. M. Lehninger Principles of Biochemistry. 5th Edition. New York: W. H.
Freeman. 2008. 1158 p.
[4] Wilson, K. & Walker, J. Principles and Techniques of Biochemistry and Molecular Biology. 7th
Edition. Cambridge: Cambridge University Press. 2010. 744 p. ISBN: 978-0-521-73167-6.
[5] New England Biolabs. T4 RNA Ligase 1 (ssRNA Ligase). 2016. [ONLINE] Available at:
https://www.neb.com/products/m0204-t4-rna-ligase-1-ssrna-ligase. [Accessed 28 October 2016].
[6] New England Biolabs. Restriction Endonucleases: Molecular Cloning and Beyond. 2016. [ONLINE]
Available at: https://www.neb.com/products/restriction-endonucleases/restrictionendonucleases/restriction-endonucleases-molecular-cloning-and-beyond. [Accessed 28 October 2016].
[7] New England Biolabs. Alkaline Phosphatase, Calf Intestinal (CIP). 2016. [ONLINE] Available at:
https://www.neb.com/products/m0290-alkaline-phosphatase-calf-intestinal-cip. [Accessed 28 October
2016].
[8] Greene, James J and Venigalla B Rao. Recombinant DNA Principles And Methodologies. New York:
Marcel Dekker, 1998. E-book. [Accessed 28 October 2016].
[9] Thermo Fisher Scientific. The Technology Behind TOPO Cloning. 2016. [ONLINE] Available at:
http://www.thermofisher.com/fi/en/home/life-science/cloning/topo/topo-resources/the-technologybehind-topo-cloning.html. [Accessed 28 October 2016].
[10] Bauman, Robert W. Microbiology With Diseases By Body System. 4th Edition. BenjaminCummings, 2015. Chapter 7. E-book.
[11] Frey, Alexander. Ligation And Transformation. 2016. Presentation.
[12] New England Biolabs.
T4 DNA Ligases. 2016. [ONLINE]
https://www.neb.com/products/m0202-t4-dna-ligase. [Accessed 28.10.2016].
Available
at:
[13] New England Biolabs. E. coli DNA Ligase. 2016. [ONLINE]
https://www.neb.com/products/m0205-e-coli-dna-ligase. [Accessed 28 October 2016]
Available
at:
5
[14] Alberts B. et al. Molecular Biology of The Cell. 6th Edition. New York: Garland Science. 2015. 1342 p. ISBN: 978-­‐0-­‐8153-­‐4464-­‐3.
6
STUDENTS: DWAMENAH RICHARD KWASI and YIN YIN
PRESENTATION DATE: 17th October, 2016
REPORT DATE: 31st October,2016
REPORT ON GEL ELETROPHORESIS
First and foremost, Gel electrophoresis basically means migrating a material through a gel
with the help of electrical current.
The first thing to do with regards to gel electrophoresis is to setup the gel matrix. Agarose is
used to separate DNA molecules and acrilamide is used to separate proteins. The gel begins
as a liquid which then is poured into a tray (molding). A comb is placed in the liquid matrix
so that when the matrix solidifies, wells are formed to load samples in them.
Immediately the Gel solidifies, it is removed from the mold and placed in a special apparatus
where electrical current is applied.
A buffer to act as an electric conductor is poured around the matrix. The samples of the
biomolecules are usually mixed with a substance of high density (viscous dye) so that they
sink to the bottom of the well instead of floating away in the buffer. The dye also helps track
the progress of the experiment. Each sample is loaded in a separate well. One of the wells is
usually assigned for loading a marker, which has a set of fragments whose sizes are already
known in order to allow for comparison with the samples being loaded.
When the current is switched on, the samples tend to move towards the positively charged
side of the apparatus since the phosphate backbones of the molecules confer a negative
charge on them. After the samples have run a sufficient distance, the matrix is studied to
view the bands that are formed by the separation of molecules.
Another electrophoresis method is SDS-PAGE. SDS-PAGE is the most commonly used to
separate proteins for purpose of analysis and purification. The generally workflow of SDSPAGE includes putting gels in the tank, filling the tank with running buffer, loading the
samples and protein standard in the gel, programming the power supply, and start to
running.
Sodium dodecyl sulfate (SDS) is a denaturing agent, which can denature secondary and nondisulfide-linked tertiary structures of proteins. Beta-mercaptoethanol (β-ME) and
dithiothreitol(DTT) are reducing agents, which can break the disulfide bridge. According to
whether added SDS, β-ME or DTT, it can be classified into native PAGE, SDS-PAGE in
reducing conditions and SDS-PAGE in non-reducing conditions. Native PAGE is the original
mode of electrophoresis. There has no breaking quaternary structure nor breaking disulfide
bond, and the entire protein stays in tact, so it allows analysis the all four levels of
biomolecular structure. When SDS-PAGE is under the non-reducing conditions, it means that
after added denaturing agents and without a reducing agent. Therefore the quaternary
structure is broken, while disulfide bonds stay in tact. Because of the utilize of antibodies in
some downstream applications of SDS-PAGE, certain antibodies only recognize protein in its
non-reduced form and the reducing agents β-mercaptoethanol and DTT must be left out of
the loading buffer and migration buffer. Another one is reducing SDS-PAGE, which is added
denaturing agents and reducing agents so that both quaternary structure and disulfide
bonds are broken. This method is widely used.
Normally the samples are intact proteins, they are single-subunit protein, or proteins with
two subunits joined by a disulfide bridge. When the samples are heated with SDS and betamercaptoethanol, SDS denatures secondary and non-disulfide-linked tertiary structures,
while beta-mercaptoethanol break the disulfide bridge between two subunits. Under
conditions that disrupt the natural structure of the sample, causing it to an unfolded linear
chain and negatively charged. Because SDS is negatively charged, it masks the intrinsic
charge of the protein. As a result, the rate at which SDS-bound protein migrates in a gel
depends primarily on its size, enabling molecular weight estimation.
For continuous systems, it means using the same buffer in the gel, sample, and electrode
reservoirs. They are used mostly for nucleic acid analysis. While discontinuous systems use a
gel separated in two sections (a large-pore stacking gel on top of a small-pore resolving gel)
and different buffers in the gels and electrode solutions.
After electrophoresis, there are so many staining methods for protein. The most common
method is Coomassie dye stains. It is especially convenient. It effectively stains proteins
within one hour, and requires only water for destaining. Silver staining is the most sensitive
colorimetric method for detecting total protein. The classical Commassie Brilliant Blue
staining can usually detect a 50ng protein bond, silver staining increases the sensitivity
typically 50 times. Unlike all other staining methods, instead of staining the proteins, Zinc
staining stains all areas of the polyacrylamide gel in which there are no proteins. Zinc
staining is as sensitive as typical silver stains (detects less than 1ng of protein).
CHEM-E8110 - Laboratory Course in Biosystems and Biomaterials Engineering
Samuli Koivu & Emmi Sveholm
Immunological detection methods
Immunological detection methods are designed to measure the presence or quantity of specific
molecules, often proteins, in biological samples. They use antibodies as probes and the detection
is based on interactions between antibodies and antigens. Different antibodies have an affinity to
bind to different molecules. This way the target molecule can be found in the sample. However,
antibodies are proteins as well, so they need to be linked to molecular labels in order to be
detected. Those labels can be for example enzymes, fluorescent dyes or radioactive chemicals.
ELISA
ELISA is short for enzyme-linked immunosorbent assay. The basic principle is that antigens are
immobilized into a solid surface, normally to a 96-well plate. A specific antibody is then applied
and it binds to the antigen. The assay uses an enzyme label that is attached to the antibody. After
this a substrate specific for the enzyme is added. This creates a reaction that produces a colorful
product which enables detection. Spectrophotometer can be used to measure the amount of the
product. There are different formats in which ELISA can be done.
Sandwich ELISA
Sandwich ELISA is the most sensitive and robust format. The name “sandwich” was given to this
format because the antigen that is measured is bound between two primary antibodies. The
procedure can be seen in picture 1. The immobilization of the antigen is accomplished by first
coating the plate with antibodies that have affinity for the antigens of interest. These antibodies
are called the capture antibodies. With this approach it is possible to bind only the wanted
antigens to the surface of the plate and all the other molecules can be washed away. After coating
the plate, the sample is added and if there is any antigen of interest present in the sample, it will
bind to the capture antibodies. The detection is accomplished by adding detector antibodies to the
plate. The detector antibodies have affinity for the antigen but they also have the enzyme label
attached to them.
Picture 1. Sandwich ELISA.
Other ELISA formats
Picture 2 presents the most popular ELISA formats. The word “direct” or “indirect” can either refer
to the capture strategy or to the detection strategy.
Picture 2. The most popular ELISA formats. The picture also lists whether the format is direct or
indirect concerning the capture and detection strategies.
In direct capture the antigen is directly immobilized to the surface of the plate instead of coating
the plate with capture antibodies like in the indirect strategy. It is not a specific method so all the
components can be absorbed to the surface of the plate, not only the antigens we want.
Like mentioned earlier, also the detection strategy can be either direct or indirect. In the direct
assay, the enzyme used for detection is directly bound to the antibody that binds to the antigen,
so the primary antibody is labeled. In the indirect assay, a labeled secondary antibody is used and
the secondary antibody binds to the primary antibody that has no label.
Sensitivity and specificity
The detection step largely determines the sensitivity of an ELISA. The indirect detection strategy is
the most popular one. It is more versatile because the same labeled secondary antibody can be
used for many primary antibodies compared to the direct detection in which all the different
primary antibodies need to be labeled for each specific ELISA system. In the indirect detection the
maximum immunoreactivity is reached for the primary antibody because it is not labeled and the
sensitivity is increased because each primary antibody has several epitopes where the labeled
secondary antibody can bind to. However, in sandwich ELISA the secondary antibody should be
specific for the detection antibody only and not for the capture antibody or the assay will not be
specific for the antigen. This can be achieved by using antibodies from different species as the
capture and primary antibodies e.g. mouse and rabbit IgG.
It is also important that the used ELISA system is specific for the molecule of interest. Crossreactivity can cause false positive results or affect the quantitation. Specificity can be tested by
verifying that the molecule of interest is detected without cross-reactivity with other closely
related molecules. For example the cross-reactivity for mouse IL-6 can be tested with rat IL-6 or
mouse IL-4.
SDS-PAGE and immunoblotting
Another common immunological detection method is to use SDS-PAGE (Sodium Dodecyl Sulfate –
Polyacrylamide Gel Electrophoresis) and immunoblotting to analyze samples. SDS is a negatively
charged molecule, which sticks to the protein of interest and keeps it denatured while it goes
through the gel electrophoresis, which separates the proteins based on their size. The proteins are
added to the gel and a current is run through it, which starts to move the proteins that are coated
in the negatively charged SDS (picture 3).
Picture 3. Protein separation using SDS-PAGE.
Immunoblotting
After SDS-PAGE is finished, the gel is removed and added on top of a nitrocellulose or
polyvinylidene difluoride (PVDF) membrane. A current is run through the gel and membrane,
which moves the negatively coated proteins from the gel to the membrane. These membranes
bind proteins very well, so before final analysis the membrane needs to be blocked to prevent
unwanted proteins (or in this case, antibodies) from binding to it. This is done by using a cheap
protein such as Bovine serum albumin (BSA). The protein in question binds to all unbound parts of
the membrane, thus blocking further binding by other proteins. After this comes the last step,
which follows steps similar to the ELISA method. Commonly a primary and secondary antibody are
used. As with ELISA, primary antibody is specific to the protein of interest and binds to it, while
secondary antibody has the signaling part and binds to the primary antibody. The different phases
of immunoblotting are shown in picture 4.
Picture 4. Immunoblotting.
In ELISA, native proteins are used but in immunoblotting the proteins are denatured. Therefore,
not all antibodies work for both techniques. This results from the fact that native proteins can
have epitopes that consist of parts that are from different loops of the protein chain so when the
protein is denatured, these parts are no longer close to each other and the epitope disappears.
Substrates and detection
There are several different methods of detection that can be used with the secondary (or in some
cases primary) antibody. One of the most common method is to use HRP (Horseradish
peroxidase). HRP is an enzyme which can be used with different types of substrates to produce
different detectable signals. With chromogenic(/colorimetric) substrates, HRP converts the
substrates to colored product molecules and the color can be seen with either eyes or special
equipment. With chemiluminescent substrates, light is produced when substrate is converted into
the product. Other possible usable methods of detection include fluorescence. These are divided
into 2 categories, where it is possible to either have the fluorescent molecule either directly
attached to the secondary antibody, or it’s possible to use an enzyme, which converts a used
substrate into a fluorescent product.
Picture 5. Different substrates and detection methods.
Radioimmunoassay
Another immunological detection method is radioimmunoassay. In this method, radioactive
substances are used for detection together with the antibodies. This method is very specific, but
the use of radioactive substances limits its usage, as it requires special equipment and usage of
radioactive substances has its own risks and requires special licensing.
Immunofluorescence
Immunofluorescence is a technique in which a specific target is labeled with a fluorescent dye
using an antibody. Samples can be for example thin sections of tissue or whole cells that have
been isolated from tissue or cultured in the lab. After staining, the sample can be examined under
a microscope. This allows the visualization of the target molecule’s distribution in the sample. In
immunofluorescence it is possible to use both primary and secondary antibodies as well, so the
detection can be direct or indirect.
CHEM-E8110 Laboratory Course in Biosystems and Biomaterial Engineering
Group II. Responsible writers: Eero & Laura
Report on presentation: Protein purification
This report is written based on the presentation given during the course. The presentation
covered basic methods in protein purification. The methods discussed are ion-exchange
chromatography, gel-filtration chromatography, affinity chromatography, pull-down assay,
and immunoprecipitation. Pull-down assay and immunoprecipitation, discussed last, are
types of affinity chromatography but are here discussed as small scale assays whereas the
three methods discussed first are column chromatographies. Other types of protein
purification include covalent chromatography and hydrophobic interaction chromatography.
In column chromatography, a sample containing the protein of interest is applied on top of a
column. Next, a solvent is added continuously, carrying the proteins through the column. The
column consists of beads that interact differently with different proteins due to the properties
of the protein. For example in ion-exchange chromatography, the beads have either a
positive or a negative charge. Thus they attract proteins with a charge of the opposite sign
and slow them down in the column. The other proteins flow through faster and in this way
different proteins are separated (Figure 1).
Figure 1​. The sample is applied to the column and carried through by using a solvent. The beads of the column
interact with proteins slowing them down. The more proteins interact the slower they move through the column.
Since different proteins interact differently proteins are separated from each other.
As mentioned above, in ion-exchange chromatography the beads have either negative or
positive charge and they attract proteins with a charge of the opposite charge. These
proteins are retarded because they spend more time in stationary phase (bound to the
beads) in comparison to other proteins which stay in mobile phase (moving and not bound)
all the way through [1] (Figure 2). Consequently, in ion-exchange chromatography the
proteins are separated based on their charge. The beads can also contain both negative and
positive groups [1]. These beads are said to be dipolar [1].
The protein binding to the beads can be manipulated by adjusting pH. At pH values that are
far from the pI of the protein, the binding is strong, and vice versa. When pH is set to the pI
value the net charge of the protein is neutral and it does not bind to a bead at all. Therefore,
a solution with pH set to pI can be used to elute proteins, which previously have attached to
beads. Another way to elute the attached proteins is to use a salt gradient (Figure 3). The
competing ions from the salt bind to the beads, thus replacing the proteins. The proteins with
the weakest interaction are replaced first, after which salt concentration is increased and
proteins with a stronger interaction are released from the column. [1]
Figure 2. ​In ion-exchange chromatography
the beads have a charge and they bind to
the proteins with a charge of the opposite
sign. Other proteins flow past the beads.
Figure 3. ​In low salt concentrations the unbound proteins flow
through the ion-exchange column. The proteins bound by the
beads can be eluted by gradually increasing the salt
concentration. The proteins with weakest interaction are
released early and the proteins with stronger interaction are
released later in in the gradient.
The gel-filtration chromatography, also known as size-exclusion chromatography, separates
proteins based on size. The beads of the column are porous. Small proteins are able to
enter the pores. Therefore, the small proteins travel a longer distance through the beads and
take more time to flow through the column in comparison to larger proteins (Figure 4).
Naturally, the pore size can be manipulated. However, unlike in ion-exchange
chromatography and in affinity chromatography, the proteins in gel-filtration chromatography
cannot be bound to the beads but are only slowed down [1]. Thus, the time has a more
significance than in ion-exchange chromatography or the affinity chromatography. The
method is convenient for example in separating multimers from their monomers [1]. This
might be difficult by using ion-exchange or affinity chromatography since mono- and
multimers might have similar electric and affinity properties. Moreover, gel-filtration
chromatography is useful tool to get rid of the salt from for example salt gradient of
ion-exchange chromatography, and for buffer change which might be needed if, for instance,
ion-exchange and affinity chromatography are done in a row and they require different
buffers.
Figure 4​. Small proteins are able to enter the porous beads in the column. Therefore they are retarded in
comparison to the larger proteins.
Affinity chromatography is based on specific interaction between ligand and target protein.
The principle of affinity chromatography is as follows. The ligand is immobilized to bead.
When protein solution is eluted through the column, proteins with specific interaction or
affinity to the ligand are bound to the ligand whereas the rest of the protein are simply eluted
out of the column. After this the target protein could be unbind from the ligands and eluted
out of the column, possibly very highly purificated.
The affinity chromatography is indeed the most specific and efficient of protein purification
methods discussed in this presentation. This is due to the fact that it relies on the specific
interactions i.e. only the target protein has the specific affinity to the ligand. In contrast,
ion-exchange and gel-filtration are based on non-specific interactions. This could be very
powerful feature especially if the protein solution consists of many different proteins or is
complex in an other way. [1]
One type of affinity chromatography or purification is a pull-down assay. It has a so called
bait-protein as a ligand, which is used to bind the target or prey protein (Figure 5). The bait
protein is modified so that it binds efficiently only to the target protein whereas the other
proteins do not interact with the bait. The bait itself has a specific tag. The tag allows it to
bind to the bead, which leads to immobilization of the bait protein. After immobilization the
bait is incubated with the protein solution, during which the prey protein is bind to the bait
protein. Then unbound proteins are washed away and, finally, the prey protein or the
bait-prey-complex can be eluted off from the bead. Pull-down assay is a great method to
research protein interactions and for protein purification. An example of pull-down assay is
described in figure 5 which shows purification of immunoglobulin by immobilized protein A/G.
Additionally, immobilizing of immunoglobulin by protein A/G can be used as method to attach
immunoglobulin to bead for acting as ligand in immunoprecipitation. [2]
Figure 5. ​Protein A/G is immobilized to agarose bead. Protein A/G has specific ability to bind with
immunoglobulin. The unbound proteins are washed out and, finally, immunoglobulin is detached from protein A/G
resulting in extremely pure immunoglobulin. [2]
Immunoprecipitation is another form of affinity purification. Immunoprecipitation is almost
identical to pull-down assay but it always uses antibody as a ligand. Immunoprecipitation is
useful method to purify or isolate antigens. The principle of immunoprecipitation is presented
in Figure 6.
Figure 6. The principle of immunoprecipitation. First the antibody is immobilized to bead after which the sample
containing antigen is added. The antigen binds to antibody forming immune complex which is eluted off the bead
and unbind later on. [2]
References
1. Janson, Jan-Christer. Methods of Biochemical Analysis : Protein Purification :
Principles, High Resolution Methods, and Applications (3). Hoboken, US: Wiley,
2011. ProQuest ebrary. Web. 27 October 2016.
2. ThermoFisher Scientific. Webpage reviewed 27 October 2016.
https://www.thermofisher.com/fi/en/home/life-science/protein-biology/protein-biology-l
earning-center/protein-biology-resource-library/pierce-protein-methods.html