Quantitative Estimation of DNA Concentrations

DNA, RNA, and protein strongly absorb ultraviolet light in the 260 to 280 nm range. UV spectroscopy can be used as a quantitative technique to measure nucleic acid concentration and protein contamination. Nucleic acids strongly absorb at 260 nm and less strongly at 280 nm while proteins do the opposite. The general rules for determining the concentrations of nucleic acids at 260 nm are:

1. 1 Optical Density (OD) unit of double-stranded DNA is 50 micrograms/ml.
2. 1 OD unit of single-stranded DNA is 33 micrograms/ml.
3. 1 OD unit of single-stranded RNA is 40 micrograms/ml.

Proteins absorb strongly at 280 nm where 1 OD unit is 1 mg/ml. When using UV spectroscopy for estimating DNA concentrations, it is very important to remove all protein and RNA from the DNA solution. Good estimations can only be made on clean preparations.

An estimate of the purity of a DNA preparation can be made by measuring the absorbance at both 260 nm and 280 nm. Pure solutions of nucleic acid will absorb approximately twice as much at 260 nm as at 280 nm. Experimentally, the ratio of 260 nm/280 nm of a pure DNA solution is between 1.8 to 2.0. As protein contamination increases, the ratio decreases. Additionally, the presence of contaminating oranic solvents, such as phenol, can affect estimations of concentration and purity.

Materials you need are:

UV Spectrophotometer

Quartz or UV compatible cuvettes

TE buffer

DNA template

Method:

1. Fill the cuvette with water or TE buffer. Zero the spectrophotometer at 260 nm with this blank.
2. DNA from plasmid and genomic preparations is typically at a concentration exceeding 1 micrograms/microliter. Consequently, DNA is usually diluted before measuring its absorbance. An unfortunate result of this measurement is that the DNA is expended as a result of the dilution. Be sure these is adequate DNA to waste. Start by diluting the DNA sample 1 microliter : 999 microliters of TE buffer (the dilution can be done directly in the cuvette). Mix the dilution thoroughly.
3. Measure the optical density (OD). Multiply the resulting OD by 50 micrograms/ml. For a 1:1000 dilution, the mass of DNA is equal to micrograms/microliter.
4. Similarly, the same sample can be measured at 280 nm. A ratio of the OD-260nm/OD-280nm is an indicator of DNA purity. A ratio of 1.8 or higher indicates minimal protein contamination.

Isolation Of Genomic DNA Rrom Yeast

Isolating genomic DNA from yeast involves culturing the microbe, harvesting the cell, enzymatically removing the cell wall, lysing the protoplast, and finally separating the DNA from the other cell debris.
Materials that you need are:

Yeast culture-prepared previously

Spectrophotometer with cuvettes

50 mM EDTA, pH 8-ice cold

50 mM Tris, pH 9.5, 2% 2-mercaptoethanol

1.2 M sorbitol, 50 mM Tris, pH 7.5

Lyticase solution-500 U/ml in 50 mM Tris, pH 7.5

10% Sodium Dodecyl Sulfate (SDS)-used for checking protoplast formation

Lysis buffer-100 mM Tris, pH 7.5, 100 mM EDTA, 150 mMNaCl, 50 micrograms/ml RNase A

Lysis buffer with 2% SDS

95% Ethanol-stored at minus 20 degree Celcius

TE buffer-10 M Tris, pH 8, 1 mM EDTA

3 M potassium acetate, pH 5.5

Here are the step by step methods:

1. The yeast can be cultured for as long as 48 hours at 30 degree Celcius. The optical density of a 1:10 dilution of the culture in water can be as high as 1.0 at 520 nm.
2. Harvest 5 ml of cells by centrifugation (5 minutes at 5000 rpm). Resuspend the yeast in 1 ml of cold 50 mM EDTA, pH 8, and transfer to a 1.5 ml microfuge tube. Centrifuge for 1 minute, decant, and resuspend again in 50 mM EDTA.
3. Pellet the cells as before and suspend the cells in 1 ml of 50 mM Tris, pH 9.5, 2% 2-mercaptoethanol. Incubate for 10 min at room temperature. Centrifuge and decant.
4. Resuspend the cells in 800 micro liter of 1.2 M sorbitol, 50 mM Tris, pH 7.5. The sorbitol act as an osmotic support and prevents rupture of the cells as the wall is removed. As the yeast cell walls degrade, membranes can easily overextend and rupture.
5. Add 200 micro liter of Lyticase (500 U/ml in 50 mM Tris, pH 7.5). Place the cells on a rocker and incubate at 37 degree Celcius for one hour. Lyticase is a yeast cell wall degrading enzyme isolated from the bacteria Arthrobacter luteus.
6. Examine the suspension under a microscope to ensure protoplast formation. As the yeast wall is degraded, the cell membrane can ooze out of the sack. Viewed with phase contrast microscopy, yeast protoplasts are characteristically refractile (or bright) spheres, and yeast cell wall shells appear as gray ghosts (cell walls without membrane and cytosol). Combine 10 micro liter of 10% SDS with 10 micro liter of yeast protoplasts. Examine the cells under the microscope. The absence of refractile yeast indicates the protoplasts were lysed by the SDS.
7. Pellet the protoplasts by centrifuging at 10000 rpm for five minutes. Resuspend the cells in 1 ml of 100 mM Tris, pH 7.5, 100 mM EDTA, 150 mM NaCl (lysis buffer). Transfer the cells to a 5 ml polypropylene tube. Add 1 ml of lysis buffer with 2% SDS. Mix and incubate at 30 Celcius egree for 30 minutes. Check the cells under a microscope for lysis.
8. Centrifuge the lysate at 5000 rpm for 15 min to pellet cellular debris. Decant the upper phase containing the DNA.
9. Using a pipet, determine the volume of the DNA solution. Add 1/10th volume (e.g., 100 micro liter for every ml) of 3 M potassium acetate to the solution. In the presence of ions Na and K, DNA precipitates if mixed with either ethanol or isopropanol. Incubate the DNA at -20 Celcius degree for 30 min (or overnight if possible. Centrifuge the solution at 7000 rpm for 20 minutes. The DNA appears as white pellet. Decant and remove as much moisture as possible, but do not allow the pellet to dry. Once genomic DNA drys, it can be very difficult to resuspend.
10. Resuspend the DNA in 100 micro liter of TE buffer and freeze.

Gold Nanoparticles In Cancer Cell Detection

“Gold nanoparticles are very good at scattering and absorbing light,” said Mostafa El-Sayed, director of the Laser Dyanamics Laboratory and chemistry professor at Georgia Tech. “We wanted to see if we could harness that scattering property in a living cell to make cancer detection easier. So far, the results are extremely promising.”

Many cancer cells have a protein, known as Epidermal Growth Factor Receptor (EFGR), all over their surface, while healthy cells typically do not express the protein as strongly. By conjugating, or binding, the gold nanoparticles to an antibody for EFGR, suitably named anti-EFGR, researchers were able to get the nanoparticles to attach themselves to the cancer cells.

“If you add this conjugated nanoparticle solution to healthy cells and cancerous cells and you look at the image, you can tell with a simple microscope that the whole cancer cell is shining,” said El-Sayed. “The healthy cell doesn’t bind to the nanoparticles specifically, so you don’t see where the cells are. With this technique, if you see a well defined cell glowing, that’s cancer.”

In the study, researchers found that the gold nanoparticles have 600 percent greater affinity for cancer cells than for noncancerous cells. The particles that worked the best were 35 nanometers in size. Researchers tested their technique using cell cultures of two different types of oral cancer and one nonmalignant cell line. The shape of the strong absorption spectrum of the gold nanoparticles are also found to distinguish between cancer cells and noncancerous cells.

What makes this technique so promising, said El-Sayed, is that it doesn’t require expensive high-powered microscopes or lasers to view the results, as other techniques require. All it takes is a simple, inexpensive microscope and white light.

Another benefit is that the results are instantaneous. “If you take cells from a cancer stricken tissue and spray them with these gold nanoparticles that have this antibody you can see the results immediately. The scattering is so strong that you can detect a single particle,” said El-Sayed.

Finally, the technique isn’t toxic to human cells. A similar technique using artificial atoms known as Quantum Dots uses semiconductor crystals to mark cancer cells, but the semiconductor material is potentially toxic to the cells and humans.

Human Development and Stem Cells

Human embryonic development depends on stem cells. During the course of development, cells divide, migrate, and specialize. Early in development, a group of cells called the inner cell mass (ICM) forms. These cells are able to produce all the tissues of the body. Later in development, during gastrulation, the three germ layers form, and most cells become more restricted in the types of cells that they can produce.

Real-time PCR in Microbiology: From Diagnosis to Characterization

Ian M. MacKay "Real-time PCR in Microbiology: From Diagnosis to Characterization"

Publisher: Caister Academic Press 2007 | 454 Pages | ISBN: 1904455182 | PDF | 15.8 MB
Description or mention of instrumentation, sofhvare, or other products in this book does not imply endorsement by the author or publisher. The author and publisher do not assume responsibility for the validity of any products or procedures mentioned or described in this book or for the consequences of
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Scripps Research Scientists Find E. Coli Enzyme Must Move to Function

Scripps Research Scientists Find E. Coli Enzyme Must Move to Function

Slight oscillations lasting just milliseconds have a huge impact on an enzyme's function, according to a new study by Scripps Research Institute scientists. Blocking these movements, without changing the enzyme's overall structure or any of its other properties, renders the enzyme defective in carrying out chemical reactions.

The study, published in the April 8, 2011 issue of the journal Science, adds to a growing body of evidence pointing to the importance of movement in the ability of enzymes and other types of proteins to do their job. The findings may also help scientists design more specific and effective drugs targeting enzymes.

"Ever since the first X-ray structures of proteins emerged, scientists have been talking about proteins as though their structures were fixed in space," said Peter Wright, chair of the Department of Molecular Biology and member of the Skaggs Institute for Chemical Biology at Scripps Research, who was senior author of the study. "But that is not how proteins work. They are like the machines we build. They have moving parts, and they need motion to work."

A Model Enzyme

The new study examined the enzyme dihydrofolate reductase (DHFR) from the common bacterium Escherichia coli, which the Wright group has been using as a model for understanding how enzymes catalyze (cause or accelerate) chemical reactions. Most strains of E. coli are harmless, but some can cause serious food poisoning.

Bacterial cells cannot live without DHFR, thus this enzyme is the target of many antibiotics. Human cells, and in particular rapidly dividing cells, also use DHFR; drugs that target human DHFR, such as methotrexate, are often used in cancer chemotherapy.

DHFR spurs the conversion of a compound called dihydrofolate (DHF) to a different form, tetrahydrofolate (THF), which is needed by cells for synthesis of DNA. In its chemical reaction, DHFR uses a helper or co-factor, called NADPH. It catalyzes the transfer of a hydride (a negative hydrogen ion) from NADPH to DHF to produce THF. Previous studies by Wright and others have shown that the loops surrounding the active site are flexible, and that one of the loops in particular, called the Met20 loop can adopt two different conformations during the catalytic cycle.

Until now, however, the significance of these motions remained obscure.

Linking Motion to Function

Wright, graduate student Gira Bhabha, and colleagues from both Scripps Research and Pennsylvania State University decided to investigate.

For the new study, the scientists turned to an imaging technique known as nuclear magnetic resonance (NMR) spectroscopy, in combination with X-ray crystallography. Unlike X-ray crystallography, a technique used to determine the structure of proteins in crystals, recently developed NMR methods allow scientists to visualize the motions of proteins in solution. The technique can capture protein motions "in a time scale that is relevant to biology, from microseconds to milliseconds to seconds," said Wright.

To determine the importance of the oscillations, the team set out to make a mutation in the DHFR enzyme that prevented the flexible Met20 loop from moving. To know which amino acids to change, the scientists compared the bacterial DHFR protein sequence to that of the human enzyme, since in the human enzyme, the Met20 loop is more rigid.

Using this approach, the scientists successfully produced a rigidified mutatant E. coli DHFR. When the scientists examined it using X-ray crystallography, they could see the mutant enzyme's structure was almost identical to the wild-type enzyme. However, NMR analysis revealed that the Met20 loop and other parts of the active site were no longer flexible in the mutant.

Significantly, the mutated E. coli enzyme transferred hydride at a rate that was 16-fold slower than that of the wild-type enzyme—a substantial loss in enzyme function.

"We demonstrated that locking down the motion in the active site prevents catalysis," said Wright.

While previous work had indicated that enzymes can exist in different shapes and forms and that changes in enzyme shape enable enzymes to bind to their substrates and co-factors or release the products, "This is the first demonstration that motions play a role in the actual chemistry of a reaction," said Wright.

Clamping Down on the Active Site

The scientists reason that when the E. coli DHFR carries out its chemical reaction, motions in the active site assist in pushing NADPH and DHF closer to one another. This proximity makes the transfer of the hydride from NAPDH to DHF more efficient. If the active site can't move, the molecules are not sufficiently close to one another for the chemical reaction to occur. "We think that the mutations prevent the enzyme from clamping down on the hydride donor and acceptor, so they can no longer get as close to each other as is necessary for efficient catalysis," explained Bhabha.

Taking motion into account when designing drugs to either inhibit or increase enzyme function could result in more effective or more specific drugs. For example, because the motions in the bacterial DHFR differ from those in the human enzyme, this difference might be exploited to design drugs that are specific for the bacterial enzyme. "It might help reduce the serious side effects of drugs that target DHFR," said Wright.

"The idea is to harness these motions in drug design," added Bhabha. "It's a difficult and challenging problem, but it could have huge impact."

In addition to Wright and Bhabha, co-authors for the paper "A dynamic knockout reveals that conformational fluctuations influence the chemical step of enzyme catalysis," include Damian C. Ekiert, Ian A. Wilson, and H. Jane Dyson at Scripps Research, and Jeeyeon Lee, Jongsik Gam, and Stephen J. Benkovic at Pennsylvania State University.

The research was supported by the National Institutes of Health and the Skaggs Institute for Chemical Biology.

About The Scripps Research Institute

The Scripps Research Institute is one of the world's largest independent, non-profit biomedical research organizations. Scripps Research is internationally recognized for its discoveries in immunology, molecular and cellular biology, chemistry, neuroscience, and vaccine development, as well as for its insights into autoimmune, cardiovascular, and infectious disease. Headquartered in La Jolla, California, the institute also includes a campus in Jupiter, Florida, where scientists focus on drug discovery and technology development in addition to basic biomedical science. Scripps Research currently employs about 3,000 scientists, staff, postdoctoral fellows, and graduate students on its two campuses. The institute's graduate program, which awards Ph.D. degrees in biology and chemistry, is ranked among the top ten such programs in the nation. For more information, see www.scripps.edu .

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