Akathisia is a movement disorder characterized by a feeling of inner restlessness and inability to stay still. Usually, the legs are most prominently affected. Those affected may fidget, rock back and forth, or pace, while some may just have an uneasy feeling in their body. The most severe cases may result in aggression, violence or suicidal thoughts.
Treatment may include switching to an antipsychotic with a lower risk of the condition. Medications with tentative evidence of benefit include diphenhydramine, trazodone, benztropine, mirtazapine, and beta blockers[DJS -- Cogentin?]. Vitamin B6 or correcting iron deficiency may also be useful. Around half of people on antipsychotics develop the condition. The term was first used by Czech neuropsychiatrist Ladislav Haškovec, who described the phenomenon in 1901. It is from Greeka-, meaning "not", and καθίζεινkathízein, meaning "to sit", or in other words an "inability to sit".
Signs and symptoms
Symptoms of akathisia may vary from a mild sense of disquiet or anxiety to a sense of terror.
People typically pace for hours because the pressure on the knees
reduces the discomfort somewhat; once their knees and legs become
fatigued and they are unable to continue pacing, they sit or lie down,
although this does not relieve the akathisia. When misdiagnosis occurs
in antipsychotic neuroleptic-induced akathisia, more antipsychotic may
be prescribed, potentially worsening the symptoms.
Neuro-psychologist Dennis Staker had drug-induced akathisia for
two days. His description of his experience was this: "It was the worst
feeling I have ever had in my entire life. I wouldn't wish it on my
worst enemy." Many patients describe symptoms of neuropathic pain akin to fibromyalgia and restless legs syndrome (RLS).
In Han et al. (2013), the authors describe restless legs syndrome's
relation to akathisia, "Some researchers regard RLS as a 'focal
akathisia' [in the legs]."
Although these side effects disappear quickly and remarkably when the
medication is stopped, tardive, or late-persisting akathisia may go on
long after the offending drug is discontinued, sometimes for a period of
years.
Healy, et al. (2006), described the following regarding akathisia: tension, insomnia, a sense of discomfort, motor restlessness, and marked anxiety and panic.
...[It comes] from so deep inside
you, you cannot locate the source of the pain … The muscles of your
jawbone go berserk, so that you bite the inside of your mouth and your
jaw locks and the pain throbs. … Your spinal column stiffens so that you
can hardly move your head or your neck and sometimes your back bends
like a bow and you cannot stand up. … You ache with restlessness, so you
feel you have to walk, to pace. And then as soon as you start pacing,
the opposite occurs to you; you must sit and rest. Back and forth, up
and down you go … you cannot get relief …
In addition, not all observable restless motion is akathisia. For
example, mania, agitated depression, and attention deficit hyperactivity
disorder may look like akathisia, but the movements feel voluntary and
not due to restlessness.
The most severe cases of akathisia have been linked to aggression, violence or suicidal ideation.
However, some reviews have noted that a link to suicide may be
confounded by the pre-existing psychiatric conditions being treated. Those with akathisia-linked suicidal ideation have tended to be younger and already more depressed or suicidal,
and it has been hypothesised that akathisia may contribute an additive
effect on top of these conditions that leads to the suicidal ideation.
Causes
Drug-induced
Akathisia is frequently associated with the use of dopamine receptor antagonistantipsychotic drugs. Understanding is still limited on the pathophysiology of akathisia, but it is seen to be associated with medications which block dopaminergic
transmission in the brain. Additionally, drugs with successful
therapeutic effects in the treatment of medication-induced akathisia
have provided additional insight into the involvement of other
transmitter systems. These include benzodiazepines, β-adrenergic blockers, and serotonin antagonists. Another major cause of the syndrome is the withdrawal observed in drug-dependent
individuals. Since dopamine deficiency (or disruptions in dopamine
signalling) appears to play an important role in the development of RLS,
a form of akathisia focused in the legs,
the sudden withdrawal or rapidly decreased dosage of drugs which
increase dopamine signalling may create similar deficits of the chemical
which mimic dopamine antagonism and thus can precipitate RLS. This is
why sudden cessation of opioids, cocaine, serotonergics, and other
euphoria-inducing substances commonly produce RLS as a side-effect.
It has been correlated with Parkinson's disease and related syndromes. It is unclear, however, whether this is due more to Parkinson's or the drugs used to treat it, such as carbidopa/levodopa (levocarb).
Antidepressants can also induce the appearance of akathisia, due to increased serotonin signalling within the central nervous system. This also explains why serotonin antagonists are often a very effective treatment.
The 2006 UK study by Healy et al. observed that akathisia is often miscoded in antidepressant clinical trials as "agitation, emotional lability, and hyperkinesis (overactivity)". The study further points out that misdiagnosis of akathisia as simple motor restlessness occurs, but that this is more properly classed as dyskinesia.
It was discovered that akathisia involves increased levels of the neurotransmitter norepinephrine, which is associated with mechanisms that regulate aggression, alertness, and arousal.
The table below summarizes factors that can induce akathisia, grouped by type, with examples or brief explanations for each:
The presence and severity of akathisia can be measured using the Barnes Akathisia Scale, which assesses both objective and subjective criteria.
Precise assessment of akathisia is problematic, as it is difficult to
differentiate from a multitude of disorders with similar symptoms. In a
study of movement disorders induced by neuroleptics, akathisia was found in only 26% of patients originally diagnosed with akathisia.
The primary distinguishing features of akathisia in comparison with
other syndromes are primarily subjective characteristics, such as the
feeling of inner restlessness. Akathisia can commonly be mistaken for agitation secondary to psychotic symptoms or mood disorder, antipsychotic dysphoria, restless legs syndrome (RLS), anxiety, insomnia, drug withdrawal states, tardive dyskinesia, or other neurological and medical conditions.
Additionally, the controversial diagnosis of "pseudoakathisia" is
given, as noted by Mark J. Garcia. In his article discussing akathisia
among adults with severe and profound intellectual disability, he
describes pseudoakathisia as "comprising all the symptoms of abnormal
movements seen with akathisia, but without a sense of restlessness".
Classification
Acute akathisia
Duration of less than 6 months
Develops soon after:
Starting antipsychotic medication or following dose increase
Probably due to neurotoxicity of antidopaminergic drugs
Withdrawal or "rebound" akathisia
Associated with switching antipsychotic medications
Onset usually within 6 weeks of discontinuation or dose decrease
Anticholinergic discontinuation reaction
Treatment
Case reports and small randomized studies suggest benzodiazepines, propranolol, and anticholinergics may help treat acute akathisia, but are much less effective in treating chronic akathisia. Taylor et al. found success in lowering the dose of antipsychotic medication as an initial response to drug-induced akathisia, which should be done gradually, if possible. To minimize the risk of akathisia from antipsychotics, the clinician is advised to be conservative when increasing dosages.
One study showed vitamin B6 to be effective for the treatment of neuroleptic-induced akathisia.
Published epidemiological data for akathisia are mostly limited to treatment periods preceding the arrival of second-generation antipsychotics. Sachdev (1995) reported an incidence rate of acute akathisia of 31% for 100 patients treated for 2 weeks with antipsychotic medications. Sachdev (1995) reported a prevalence range from 0.1% to 41%. In all likelihood, rates of prevalence are lower for current treatment as second-generation antipsychotics carry a lower risk of akathisia.
Gel
electrophoresis apparatus – an agarose gel is placed in this
buffer-filled box and an electrical field is applied via the power
supply to the rear. The negative terminal is at the far end (black
wire), so DNA migrates toward the positively charged anode (red wire).
Digital
image of 3 plasmid restriction digests run on a 1% w/v agarose gel, 3
volt/cm, stained with ethidium bromide. The DNA size marker is a
commercial 1 kbp ladder. The position of the wells and direction of DNA
migration is noted.
The
image above shows how small DNA fragments will migrate through agarose
gel farther than large DNA fragments during electrophoresis. The graph
to the right shows the nonlinear, relationship between the size of the
DNA fragment and the distance migrated.
Gel
Electrophoresis is a process where an electric current is applied to
DNA samples creating fragments that can be used for comparison between
DNA samples. 1) DNA is extracted. 2) Isolation and amplification of DNA. 3) DNA added to the gel wells. 4) Electric current applied to the gel. 5) DNA bands are separated by size. 6) DNA bands are stained.
Gel electrophoresis is a method for separation and analysis of macromolecules (DNA, RNA and proteins)
and their fragments, based on their size and charge. It is used in
clinical chemistry to separate proteins by charge or size (IEF agarose,
essentially size independent) and in biochemistry and molecular biology
to separate a mixed population of DNA and RNA fragments by length, to
estimate the size of DNA and RNA fragments or to separate proteins by
charge.
Nucleic acid molecules are separated by applying an electric field to move the negatively charged molecules through a matrix of agarose
or other substances. Shorter molecules move faster and migrate farther
than longer ones because shorter molecules migrate more easily through
the pores of the gel. This phenomenon is called sieving.
Proteins are separated by charge in agarose because the pores of the
gel are too large to sieve proteins. Gel electrophoresis can also be
used for separation of nanoparticles.
Gel electrophoresis uses a gel as an anticonvective medium or
sieving medium during electrophoresis, the movement of a charged
particle in an electrical field. Gels suppress the thermal convection
caused by application of the electric field, and can also act as a
sieving medium, retarding the passage of molecules; gels can also simply
serve to maintain the finished separation, so that a post
electrophoresis stain can be applied. DNA Gel electrophoresis is usually performed for analytical purposes, often after amplification of DNA via polymerase chain reaction (PCR), but may be used as a preparative technique prior to use of other methods such as mass spectrometry, RFLP, PCR, cloning, DNA sequencing, or Southern blotting for further characterization.
Physical basis
Overview of Gel Electrophoresis.
Electrophoresis
is a process which enables the sorting of molecules based on size.
Using an electric field, molecules (such as DNA) can be made to move
through a gel made of agarose or polyacrylamide.
The electric field consists of a negative charge at one end which
pushes the molecules through the gel, and a positive charge at the other
end that pulls the molecules through the gel. The molecules being
sorted are dispensed into a well in the gel material. The gel is placed
in an electrophoresis chamber, which is then connected to a power
source. When the electric current is applied, the larger molecules move
more slowly through the gel while the smaller molecules move faster. The
different sized molecules form distinct bands on the gel.
The term "gel" in this instance refers to the matrix used to contain, then separate the target molecules. In most cases, the gel is a crosslinked polymer
whose composition and porosity is chosen based on the specific weight
and composition of the target to be analyzed. When separating proteins or small nucleic acids (DNA, RNA, or oligonucleotides) the gel is usually composed of different concentrations of acrylamide and a cross-linker, producing different sized mesh networks of polyacrylamide. When separating larger nucleic acids (greater than a few hundred bases),
the preferred matrix is purified agarose. In both cases, the gel forms a
solid, yet porous matrix. Acrylamide, in contrast to polyacrylamide, is
a neurotoxin
and must be handled using appropriate safety precautions to avoid
poisoning. Agarose is composed of long unbranched chains of uncharged
carbohydrate without cross links resulting in a gel with large pores
allowing for the separation of macromolecules and macromolecular complexes.
Electrophoresis refers to the electromotive force
(EMF) that is used to move the molecules through the gel matrix. By
placing the molecules in wells in the gel and applying an electric
field, the molecules will move through the matrix at different rates,
determined largely by their mass when the charge-to-mass ratio (Z) of
all species is uniform. However, when charges are not all uniform the
electrical field generated by the electrophoresis procedure will cause
the molecules to migrate differentially according to charge. Species
that are net positively charged will migrate towards the cathode which is negatively charged (because this is an electrolytic rather than galvanic cell),
whereas species that are net negatively charged will migrate towards
the positively charged anode. Mass remains a factor in the speed with
which these non-uniformly charged molecules migrate through the matrix
toward their respective electrodes.
If several samples have been loaded into adjacent wells in the
gel, they will run parallel in individual lanes. Depending on the number
of different molecules, each lane shows separation of the components
from the original mixture as one or more distinct bands, one band per
component. Incomplete separation of the components can lead to
overlapping bands, or to indistinguishable smears representing multiple
unresolved components.
Bands in different lanes that end up at the same distance from the top
contain molecules that passed through the gel with the same speed,
which usually means they are approximately the same size. There are molecular weight size markers
available that contain a mixture of molecules of known sizes. If such a
marker was run on one lane in the gel parallel to the unknown samples,
the bands observed can be compared to those of the unknown in order to
determine their size. The distance a band travels is approximately
inversely proportional to the logarithm of the size of the molecule.
There are limits to electrophoretic techniques. Since passing
current through a gel causes heating, gels may melt during
electrophoresis. Electrophoresis is performed in buffer solutions to
reduce pH changes due to the electric field, which is important because
the charge of DNA and RNA depends on pH, but running for too long can
exhaust the buffering capacity of the solution. There are also
limitations in determining the molecular weight by SDS-PAGE, especially
when trying to find the MW of an unknown protein. There are certain
biological variables that are difficult or impossible to minimize and
can affect the electrophoretic migration. Such factors include protein
structure, post-translational modifications, and amino acid composition.
For example, tropomyosin is an acidic protein that migrates abnormally
on SDS-PAGE gels. This is because the acidic residues are repelled by
the negatively charged SDS, leading to an inaccurate mass-to-charge
ratio and migration.
Further, different preparations of genetic material may not migrate
consistently with each other, for morphological or other reasons.
Types of gel
The
types of gel most typically used are agarose and polyacrylamide gels.
Each type of gel is well-suited to different types and sizes of analyte.
Polyacrylamide gels are usually used for proteins, and have very high
resolving power for small fragments of DNA (5-500 bp). Agarose gels on
the other hand have lower resolving power for DNA but have greater
range of separation, and are therefore used for DNA fragments of usually
50-20,000 bp in size, but resolution of over 6 Mb is possible with pulsed field gel electrophoresis (PFGE).
Polyacrylamide gels are run in a vertical configuration while agarose
gels are typically run horizontally in a submarine mode. They also
differ in their casting methodology, as agarose sets thermally, while
polyacrylamide forms in a chemical polymerization reaction.
Agarose
Inserting the gel comb in an agarose gel electrophoresis chamber
Agarose gels are made from the natural polysaccharidepolymers extracted from seaweed.
Agarose gels are easily cast and handled compared to other matrices,
because the gel setting is a physical rather than chemical change.
Samples are also easily recovered. After the experiment is finished, the
resulting gel can be stored in a plastic bag in a refrigerator.
Agarose gels do not have a uniform pore size, but are optimal for electrophoresis of proteins that are larger than 200 kDa. Agarose gel electrophoresis can also be used for the separation of DNA fragments ranging from 50 base pair
to several megabases (millions of bases), the largest of which require
specialized apparatus. The distance between DNA bands of different
lengths is influenced by the percent agarose in the gel, with higher
percentages requiring longer run times, sometimes days. Instead high
percentage agarose gels should be run with a pulsed field electrophoresis (PFE), or field inversion electrophoresis.
"Most agarose gels are made with between 0.7% (good separation or
resolution of large 5–10kb DNA fragments) and 2% (good resolution for
small 0.2–1kb fragments) agarose dissolved in electrophoresis buffer. Up
to 3% can be used for separating very tiny fragments but a vertical
polyacrylamide gel is more appropriate in this case. Low percentage gels
are very weak and may break when you try to lift them. High percentage
gels are often brittle and do not set evenly. 1% gels are common for
many applications."
Polyacrylamide
Polyacrylamide gel electrophoresis (PAGE) is used for separating
proteins ranging in size from 5 to 2,000 kDa due to the uniform pore
size provided by the polyacrylamide gel. Pore size is controlled by
modulating the concentrations of acrylamide and bis-acrylamide powder
used in creating a gel. Care must be used when creating this type of
gel, as acrylamide is a potent neurotoxin in its liquid and powdered
forms.
Traditional DNA sequencing techniques such as Maxam-Gilbert or Sanger
methods used polyacrylamide gels to separate DNA fragments differing by
a single base-pair in length so the sequence could be read. Most modern
DNA separation methods now use agarose gels, except for particularly
small DNA fragments. It is currently most often used in the field of immunology and protein analysis, often used to separate different proteins or isoforms of the same protein into separate bands. These can be transferred onto a nitrocellulose or PVDF membrane to be probed with antibodies and corresponding markers, such as in a western blot.
Typically resolving gels
are made in 6%, 8%, 10%, 12% or 15%. Stacking gel (5%) is poured on
top of the resolving gel and a gel comb (which forms the wells and
defines the lanes where proteins, sample buffer and ladders will be
placed) is inserted. The percentage chosen depends on the size of the
protein that one wishes to identify or probe in the sample. The smaller
the known weight, the higher the percentage that should be used. Changes
on the buffer system of the gel can help to further resolve proteins of
very small sizes.
Starch
Partially hydrolysed
potato starch makes for another non-toxic medium for protein
electrophoresis. The gels are slightly more opaque than acrylamide or
agarose. Non-denatured proteins can be separated according to charge and
size. They are visualised using Napthal Black or Amido Black staining.
Typical starch gel concentrations are 5% to 10%.
Gel conditions
Denaturing
TTGE
profiles representing the bifidobacterial diversity of fecal samples
from two healthy volunteers (A and B) before and after AMC (Oral
Amoxicillin-Clavulanic Acid) treatment
Denaturing
gels are run under conditions that disrupt the natural structure of the
analyte, causing it to unfold into a linear chain. Thus, the mobility
of each macromolecule depends only on its linear length and its mass-to-charge ratio. Thus, the secondary, tertiary, and quaternary levels of biomolecular structure are disrupted, leaving only the primary structure to be analyzed.
Nucleic acids are often denatured by including urea in the buffer, while proteins are denatured using sodium dodecyl sulfate, usually as part of the SDS-PAGE process. For full denaturation of proteins, it is also necessary to reduce the covalent disulfide bonds that stabilize their tertiary and quaternary structure, a method called reducing PAGE. Reducing conditions are usually maintained by the addition of beta-mercaptoethanol or dithiothreitol. For general analysis of protein samples, reducing PAGE is the most common form of protein electrophoresis.
Denaturing conditions are necessary for proper estimation of
molecular weight of RNA. RNA is able to form more intramolecular
interactions than DNA which may result in change of its electrophoretic mobility. Urea, DMSO and glyoxal are the most often used denaturing agents to disrupt RNA structure. Originally, highly toxic methylmercury hydroxide was often used in denaturing RNA electrophoresis, but it may be method of choice for some samples.
Denaturing gel electrophoresis is used in the DNA and RNA banding pattern-based methods temperature gradient gel electrophoresis (TGGE) and denaturing gradient gel electrophoresis (DGGE).
Native gels are run in non-denaturing conditions, so that the
analyte's natural structure is maintained. This allows the physical
size of the folded or assembled complex to affect the mobility, allowing
for analysis of all four levels of the biomolecular structure. For
biological samples, detergents are used only to the extent that they are
necessary to lyselipid membranes in the cell.
Complexes remain—for the most part—associated and folded as they would
be in the cell. One downside, however, is that complexes may not
separate cleanly or predictably, as it is difficult to predict how the
molecule's shape and size will affect its mobility. Addressing and
solving this problem is a major aim of quantitative native PAGE.
Unlike denaturing methods, native gel electrophoresis does not use a charged denaturing agent. The molecules being separated (usually proteins or nucleic acids) therefore differ not only in molecular mass
and intrinsic charge, but also the cross-sectional area, and thus
experience different electrophoretic forces dependent on the shape of
the overall structure. For proteins, since they remain in the native
state they may be visualised not only by general protein staining
reagents but also by specific enzyme-linked staining.
A specific experiment example of an application of native gel
electrophoresis is to check for enzymatic activity to verify the
presence of the enzyme in the sample during protein purification. For
example, for the protein alkaline phosphatase, the staining solution is a
mixture of 4-chloro-2-2methylbenzenediazonium salt with
3-phospho-2-naphthoic acid-2’-4’-dimethyl aniline in Tris buffer. This
stain is commercially sold as kit for staining gels. If the protein is
present, the mechanism of the reaction takes place in the following
order: it starts with the de-phosphorylation of 3-phospho-2-naphthoic
acid-2’-4’-dimethyl aniline by alkaline phosphatase (water is needed for
the reaction). The phosphate group is released and replaced by an
alcohol group from water. The electrophile 4- chloro-2-2
methylbenzenediazonium (Fast Red TR Diazonium salt) displaces the
alcohol group forming the final product Red Azo dye. As its name
implies, this is the final visible-red product of the reaction. In
undergraduate academic experimentation of protein purification, the gel
is usually ran next to commercial purified samples in order to visualize
the results and conclude whether or not purification was successful.
Buffers
in gel electrophoresis are used to provide ions that carry a current
and to maintain the pH at a relatively constant value.
These buffers have plenty of ions in them, which is necessary for the
passage of electricity through them. Something like distilled water or
benzene contains few ions, which is not ideal for the use in
electrophoresis. There are a number of buffers used for electrophoresis. The most common being, for nucleic acids Tris/Acetate/EDTA (TAE), Tris/Borate/EDTA (TBE). Many other buffers have been proposed, e.g. lithium borate,
which is almost never used, based on Pubmed citations (LB), iso
electric histidine, pK matched goods buffers, etc.; in most cases the
purported rationale is lower current (less heat) matched ion
mobilities, which leads to longer buffer life. Borate is problematic;
Borate can polymerize, or interact with cis diols such as those found in
RNA. TAE has the lowest buffering capacity but provides the best
resolution for larger DNA. This means a lower voltage and more time, but
a better product. LB is relatively new and is ineffective in resolving
fragments larger than 5 kbp; However, with its low conductivity, a much
higher voltage could be used (up to 35 V/cm), which means a shorter
analysis time for routine electrophoresis. As low as one base pair size
difference could be resolved in 3% agarose gel with an extremely low
conductivity medium (1 mM Lithium borate).
Most SDS-PAGE protein separations are performed using a "discontinuous" (or DISC) buffer system
that significantly enhances the sharpness of the bands within the gel.
During electrophoresis in a discontinuous gel system, an ion gradient is
formed in the early stage of electrophoresis that causes all of the
proteins to focus into a single sharp band in a process called isotachophoresis.
Separation of the proteins by size is achieved in the lower,
"resolving" region of the gel. The resolving gel typically has a much
smaller pore size, which leads to a sieving effect that now determines
the electrophoretic mobility of the proteins.
Visualization
After the electrophoresis is complete, the molecules in the gel can be stained to make them visible. DNA may be visualized using ethidium bromide which, when intercalated into DNA, fluoresce under ultraviolet light, while protein may be visualised using silver stain or Coomassie Brilliant Blue
dye. Other methods may also be used to visualize the separation of the
mixture's components on the gel. If the molecules to be separated
contain radioactivity, for example in a DNA sequencing gel, an autoradiogram can be recorded of the gel. Photographs can be taken of gels, often using a Gel Doc system.
Downstream processing
After separation, an additional separation method may then be used, such as isoelectric focusing or SDS-PAGE.
The gel will then be physically cut, and the protein complexes
extracted from each portion separately. Each extract may then be
analysed, such as by peptide mass fingerprinting or de novo peptide sequencing after in-gel digestion. This can provide a great deal of information about the identities of the proteins in a complex.
Applications
Estimation of the size of DNA molecules following restriction enzyme digestion, e.g. in restriction mapping of cloned DNA.
Gel electrophoresis is used in forensics, molecular biology, genetics, microbiology and biochemistry.
The results can be analyzed quantitatively by visualizing the gel with
UV light and a gel imaging device. The image is recorded with a computer
operated camera, and the intensity of the band or spot of interest is
measured and compared against standard or markers loaded on the same
gel. The measurement and analysis are mostly done with specialized
software.
Depending on the type of analysis being performed, other
techniques are often implemented in conjunction with the results of gel
electrophoresis, providing a wide range of field-specific applications.
Nucleic acids
An agarose gel of a PCR product compared to a DNA ladder.
In the case of nucleic acids, the direction of migration, from
negative to positive electrodes, is due to the naturally occurring
negative charge carried by their sugar-phosphate backbone.
Double-stranded DNA fragments naturally behave as long rods, so
their migration through the gel is relative to their size or, for cyclic
fragments, their radius of gyration. Circular DNA such as plasmids,
however, may show multiple bands, the speed of migration may depend on
whether it is relaxed or supercoiled. Single-stranded DNA or RNA tend
to fold up into molecules with complex shapes and migrate through the
gel in a complicated manner based on their tertiary structure.
Therefore, agents that disrupt the hydrogen bonds, such as sodium hydroxide or formamide, are used to denature the nucleic acids and cause them to behave as long rods again.
Gel electrophoresis of large DNA or RNA is usually done by agarose gel electrophoresis. See the "Chain termination method"
page for an example of a polyacrylamide DNA sequencing gel.
Characterization through ligand interaction of nucleic acids or
fragments may be performed by mobility shift affinity electrophoresis.
Electrophoresis of RNA samples can be used to check for genomic
DNA contamination and also for RNA degradation. RNA from eukaryotic
organisms shows distinct bands of 28s and 18s rRNA, the 28s band being
approximately twice as intense as the 18s band. Degraded RNA has less
sharply defined bands, has a smeared appearance, and intensity ratio is
less than 2:1.
Proteins
SDS-PAGE autoradiography – The indicated proteins are present in different concentrations in the two samples.
Proteins,
unlike nucleic acids, can have varying charges and complex shapes,
therefore they may not migrate into the polyacrylamide gel at similar
rates, or at all, when placing a negative to positive EMF on the sample.
Proteins therefore, are usually denatured in the presence of a detergent such as sodium dodecyl sulfate (SDS) that coats the proteins with a negative charge.
Generally, the amount of SDS bound is relative to the size of the
protein (usually 1.4g SDS per gram of protein), so that the resulting
denatured proteins have an overall negative charge, and all the proteins
have a similar charge-to-mass ratio. Since denatured proteins act like
long rods instead of having a complex tertiary shape, the rate at which
the resulting SDS coated proteins migrate in the gel is relative only to
its size and not its charge or shape.
A
novel application for the gel electrophoresis is to separate or
characterize metal or metal oxide nanoparticles (e.g. Au, Ag, ZnO, SiO2)
regarding size, shape or surface chemistry of the nanoparticles. The
scope is to obtain a more homogeneous sample (e.g. narrower particle
size distribution), which than can be used in further products/processes
(e.g. self-assemly processes). For the separation of nanoparticles
within a gel the particle size in relation to the mesh size is the key
parameter, whereby two migration mechanisms where identified: the
unrestricted mechanism, where the particle size << mesh size and
the restricted mechanism, where particle size is similar to mesh size.
History
1930s – first reports of the use of sucrose for gel electrophoresis
1955 – introduction of starch gels, mediocre separation (Smithies)
1959 – introduction of acrylamide gels; disc electrophoresis
(Ornstein and Davis); accurate control of parameters such as pore size
and stability; and (Raymond and Weintraub)
2004 – introduction of a standardized time of polymerization of acrylamide gels enables clean and predictable separation of native proteins (Kastenholz)
A 1959 book on electrophoresis by Milan Bier cites references from the 1800s. However, Oliver Smithies
made significant contributions. Bier states: "The method of Smithies
... is finding wide application because of its unique separatory power."
Taken in context, Bier clearly implies that Smithies' method is an
improvement.
In this diagram of a duplicated chromosome, (2) identifies the centromere—the region that joins the two sister chromatids, or each half of the chromosome. In prophase of mitosis, specialized regions on centromeres called kinetochores attach chromosomes to spindle fibers.
The physical role of the centromere is to act as the site of assembly of the kinetochores – a highly complex multiprotein structure that is responsible for the actual events of chromosome segregation – i.e. binding microtubules and signalling to the cell cycle machinery when all chromosomes have adopted correct attachments to the spindle, so that it is safe for cell division to proceed to completion and for cells to enter anaphase.
There are, broadly speaking, two types of centromeres. "Point centromeres" bind to specific proteins that recognize particular DNAsequences with high efficiency.
Any piece of DNA with the point centromere DNA sequence on it will
typically form a centromere if present in the appropriate species. The
best characterised point centromeres are those of the budding yeast, Saccharomyces cerevisiae.
"Regional centromeres" is the term coined to describe most centromeres,
which typically form on regions of preferred DNA sequence, but which
can form on other DNA sequences as well. The signal for formation of a regional centromere appears to be epigenetic. Most organisms, ranging from the fission yeast Schizosaccharomyces pombe to humans, have regional centromeres.
Regarding mitotic chromosome structure, centromeres represent a
constricted region of the chromosome (often referred to as the primary
constriction) where two identical sister chromatids
are most closely in contact. When cells enter mitosis, the sister
chromatids (the two copies of each chromosomal DNA molecule resulting
from DNA replication in chromatin form) are linked along their length by the action of the cohesin
complex. It is now believed that this complex is mostly released from
chromosome arms during prophase, so that by the time the chromosomes
line up at the mid-plane of the mitotic spindle (also known as the
metaphase plate), the last place where they are linked with one another
is in the chromatin in and around the centromere.
Position
Classifications of Chromosomes
I
Telocentric
Centromere placement very close to the top, p arms barely visible if visible at all.
II
Acrocentric
q arms are still much longer than the p arms, but the p arms are longer than those in telocentric.
III
Submetacentric
p and q arms are very close in length but not equal.
IV
Metacentric
p and q arms are equal in length.
A: Short arm (p arm) B: Centromere C: Long arm (q arm) D: Sister Chromatids
Each chromosome has two arms, labeled p (the shorter of the two) and q
(the longer). Many remember that the short arm 'p' is named for the
French word "petit" meaning 'small', although this explanation was shown
to be apocryphal. They can be connected in either metacentric, submetacentric, acrocentric or telocentric manner.
Categorization of chromosomes according to the relative arms length
These are X-shaped chromosomes, with the centromere in the middle so that the two arms of the chromosomes are almost equal.
A chromosome is metacentric if its two arms are roughly equal in length. In a normal human karyotype,
five chromosomes are considered metacentric: chromosomes 1, 3, 16, 19,
and 20. In some cases, a metacentric chromosome is formed by balanced
translocation: the fusion of two acrocentric chromosomes to form one metacentric chromosome.
Submetacentric
If the arms' lengths are unequal, the chromosome is said to be submetacentric. They are L-shaped.
Acrocentric
If the p (short) arm is so short that it is hard to observe, but still present, then the chromosome is acrocentric (the "acro-" in acrocentric refers to the Greek word for "peak"). The human genome includes five acrocentric chromosomes: 13, 14, 15, 21, 22. The Y chromosome is also acrocentric.
In an acrocentric chromosome the p arm contains genetic material
including repeated sequences such as nucleolar organizing regions, and
can be translocated without significant harm, as in a balanced Robertsonian translocation. The domestic horse genome includes one metacentric chromosome that is homologous to two acrocentric chromosomes in the conspecific but undomesticated Przewalski's horse.
This may reflect either fixation of a balanced Robertsonian
translocation in domestic horses or, conversely, fixation of the fission
of one metacentric chromosome into two acrocentric chromosomes in
Przewalski's horses. A similar situation exists between the human and
great ape genomes, with a reduction of two acrocentric chromosomes in
the great apes to one metacentric chromosome in humans (see aneuploidy and the human chromosome 2).
Strikingly, harmful translocations in disease context, especially
unbalanced translocations in blood cancers, more frequently involve
acrocentric chromosomes than non-acrocentric chromosomes. Although the cause is not known, this probably relates to the physical location of acrocentric chromosomes within the nucleus. Acrocentric chromosomes are usually located in and around the nucleolus, so in the center of the nucleus, where chromosomes tend to be less densely packed than chromosomes in the nuclear periphery. Consistently, chromosomal regions that are less densely packed are also more prone to chromosomal translocations in cancers.
Telocentric
A
telocentric chromosome's centromere is located at the terminal end of
the chromosome. A telocentric chromosome has therefore only one arm. Telomeres may extend from both ends of the chromosome, their shape is similar to letter "i" during anaphase. For example, the standard house mouse karyotype has only telocentric chromosomes. Humans do not possess telocentric chromosomes.
Subtelocentric
If the chromosome's centromere is located closer to its end than to its center, it may be described as subtelocentric.
Centromere number
Acentric
If a chromosome lacks a centromere, it is said acentric. The macronucleus of ciliates for example contains hundreds of acentric chromosomes. Chromosome-breaking events can also generate acentric chromosomes or acentric fragments.
Dicentric
A dicentric chromosome
is an abnormal chromosome with two centromeres. It is formed through
the fusion of two chromosome segments, each with a centromere, resulting
in the loss of acentric fragments (lacking a centromere) and the
formation of dicentric fragments. The formation of dicentric chromosomes has been attributed to genetic processes, such as Robertsonian translocation and paracentric inversion.
Dicentric chromosomes have important roles in the mitotic stability of
chromosomes and the formation of pseudodicentric chromosomes.
Monocentric
The monocentric chromosome is a chromosome that has only one centromere in a chromosome and forms a narrow constriction.
Monocentric centromeres are the most common structure on highly repetitive DNA in plants and animals.
Holocentric
Different
than monocentric chromosones in holocentric chromosomes, the entire
length of the chromosome acts as the centromere. In holocentric
chromosomes there is not one primary constriction but the centromere has
many CenH3 loci spread over the whole chromosome. Examples of this type of centromere can be found scattered throughout the plant and animal kingdoms, with the most well-known example being the nematode Caenorhabditis elegans.
Polycentric
Human chromosomes
Table of human chromosomes with data on centromeres and sizes.
There are two types of centromeres.[27] In regional centromeres, DNA
sequences contribute to but do not define function. Regional
centromeres contain large amounts of DNA and are often packaged into heterochromatin. In most eukaryotes, the centromere's DNA sequence consists of large arrays of repetitive DNA (e.g. satellite DNA)
where the sequence within individual repeat elements is similar but not
identical. In humans, the primary centromeric repeat unit is called
α-satellite (or alphoid), although a number of other sequence types are
found in this region.[28]
Point centromeres are smaller and more compact. DNA sequences are
both necessary and sufficient to specify centromere identity and
function in organisms with point centromeres. In budding yeasts, the
centromere region is relatively small (about 125 bp DNA) and contains
two highly conserved DNA sequences that serve as binding sites for
essential kinetochore proteins.[28]
Inheritance
Since centromeric DNA sequence is not the key determinant of centromeric identity in metazoans, it is thought that epigenetic inheritance plays a major role in specifying the centromere.[29]
The daughter chromosomes will assemble centromeres in the same place as
the parent chromosome, independent of sequence. It has been proposed
that histone H3 variant CENP-A (Centromere Protein A) is the epigenetic mark of the centromere.[30]
The question arises whether there must be still some original way in
which the centromere is specified, even if it is subsequently propagated
epigenetically. If the centromere is inherited epigenetically from one
generation to the next, the problem is pushed back to the origin of the
first metazoans.
Structure
The centromeric DNA is normally in a heterochromatin state, which is essential for the recruitment of the cohesin
complex that mediates sister chromatid cohesion after DNA replication
as well as coordinating sister chromatid separation during anaphase. In
this chromatin, the normal histone H3 is replaced with a centromere-specific variant, CENP-A in humans.[31]
The presence of CENP-A is believed to be important for the assembly of
the kinetochore on the centromere. CENP-C has been shown to localise
almost exclusively to these regions of CENP-A associated chromatin. In
human cells, the histones are found to be most enriched for H4K20me3 and H3K9me3[32]
which are known heterochromatic modifications. In Drosophila, Islands
of retroelements are major components of the centromeres. [33]
In the yeast Schizosaccharomyces pombe (and probably in other eukaryotes), the formation of centromeric heterochromatin is connected to RNAi.[34] In nematodes such as Caenorhabditis elegans,
some plants, and the insect orders Lepidoptera and Hemiptera,
chromosomes are "holocentric", indicating that there is not a primary
site of microtubule attachments or a primary constriction, and a
"diffuse" kinetochore assembles along the entire length of the
chromosome.
Centromeric aberrations
In rare cases in humans, neocentromeres
can form at new sites on the chromosome. There are currently over 90
known human neocentromeres identified on 20 different chromosomes.[35][36]
The formation of a neocentromere must be coupled with the inactivation
of the previous centromere, since chromosomes with two functional
centromeres (Dicentric chromosome)
will result in chromosome breakage during mitosis. In some unusual
cases human neocentromeres have been observed to form spontaneously on
fragmented chromosomes. Some of these new positions were originally
euchromatic and lack alpha satellite DNA altogether.
It
has been known that centromere misregulation contributes to
mis-segregation of chromosomes, which is strongly related to cancer and
abortion. Notably, overexpression of many centromere genes have been
linked to cancer malignant phenotypes. Overexpression of these
centromere genes can increase genomic instability in cancers.[37]
Elevated genomic instability on one hand relates to malignant
phenotypes; on the other hand, it makes the tumor cells more vulnerable
to specific adjuvant therapies such as certain chemotherapies and
radiotherapy.[38] Instability of centromere repetitive DNA was recently shown in cancer and aging.[39]
