Researchers at the University of Cincinnati (UC) are looking for ways to reduce or prevent heart damage by starting where the problem often begins: in the genes.

Following a heart attack, cells die, causing lasting damage to the heart.

Keith Jones, PhD, a researcher in the department of pharmacology and cell biophysics, and colleagues are trying to reduce post-heart attack damage by studying the way cells die in the heart—a process controlled by transcription factors.

Transcription factors are proteins that bind to specific parts of DNA and are part of a system that controls the transfer of genetic information from DNA to RNA and then to protein.  Transfer of genetic information also plays a role in controlling the cycle of cells—from cell growth to cell death.

“We call it ‘gene regulatory therapy,’” says Jones.

So far, studies have identified the role for an important group of interacting transcription factors and the genes they regulate to determine whether cells in the heart survive or die after blood flow restriction occurs.

Often, scientists use virus-like mechanisms to transfer DNA and other nucleic acids inside the body.

The “virus” takes over other healthy cells by injecting them with its DNA.  The cells, then transformed, begin reproducing the virus’ DNA.  Eventually they swell and burst, sending multiple replicas of the virus out to conquer other cells and repeat the process.

Now, UC researchers are further investigating new, non-viral delivery mechanisms for this transfer of DNA.

“We can use non-viral delivery vehicles to transfer nucleic acids, including transcription factor decoys, to repress activation of specific transcription factors in the heart,” Jones says, adding that the researchers have made this successfully work within live animal models.  “This means we can block the activity of most transcription factors in the heart without having to make genetically engineered mice.”

Jones will be presenting these results at the International Society for Heart Research in Cincinnati, June 17-20.

He says this delivery mechanism involves flooding the cells with “decoys” which trick the transcription factors into binding to the decoys rather than to target genes, preventing them from activating those genes.

“We can use this technology to identify the target genes and then investigate the action of these genes in the biological process,” Jones says.

He says that this delivery has limitations and advantages.

“It can be used to block a factor at any point in time and is reversible,” he says.  “However, right now, a specific delivery route must be used to target the tissue or cell.”

Jones and other researchers are hoping that this new technology will allow them to directly address the effects of gene regulation in disease, as opposed to using classical drugs that treat symptoms or have significant adverse outcomes.

“So far, this seems to cause no adverse effects in animals,” he says.  “We are hopeful and are working toward pre-clinical studies.”

For the first time, researchers at Delft University of Technology have witnessed the spontaneous repair of damage to DNA molecules in real time. They observed this at the level of a single DNA molecule. Insight into this type of repair mechanism is essential as errors in this process can lead to the development of cancerous cells. Researchers from the Kavli Institute of Nanoscience Delft are to publish an article on this in the leading scientific journal Molecular Cell.

Cells have mechanisms for repairing the continuous accidental damage occurring in DNA. These damages can vary from a change to a single part of the DNA to a total break in the DNA structure. These breaks can, for instance, be caused by ultraviolet light or X-rays, but also occur during cell division, when DNA molecules split and form two new DNA molecules. If this type of break is not properly repaired it can be highly dangerous to the functioning of the cell and lead to the creation of a cancerous cell.

One major DNA-repair mechanism involved in repairing these breaks is known as homologous recombination. This mechanism has been observed for the first time by Delft University of Technology researchers in real time and at the level of a single DNA molecule.

To observe this, a DNA molecule is stretched between a magnetic bead and a glass surface. A force is exerted on the magnetic bead using a magnetic field, enabling researchers to pull and rotate a single DNA molecule in a controlled fashion. As the position of the bead changes when the DNA molecule is repaired, researchers are able to observe the repair process in detail.

University of Illinois researchers have developed a technique for imaging cells under an electron microscope that yields a sharper image of the structure of chromatin, the tightly wound bundle of genetic material and proteins that makes up the chromosomes. The findings appeared in Nature Methods.

Scientists have known for more than a century that proteins, such as histones, aid in packing DNA into the nucleus of a cell.  Human cells contain 2 to 3 meters of DNA, which must be kinked and coiled enough to fit into a region 1/10 the width of a human hair.

Despite the use of powerful, high-resolution imaging techniques such as electron microscopy, the mechanism by which this chromatin packing occurs remains a mystery.  The densely coiled chromatin fibers are very difficult to visualize, and little is known about how they condense during cell division, or unwind to allow gene expression.

In developing their method, the Illinois team tackled a key difficulty in imaging cells using electron microscopy.  Traditional studies “fix” the cells with potent chemicals (called fixatives) to preserve their structure for viewing under a microscope.  But standard fixation methods interfere with another step in the imaging process: the use of tagged antibodies to label key components of the cells.

These antibodies, which target and latch on to specific proteins in the cell, can be tagged with fluorescent labels for detection in light microscopy, or with metal particles (gold, in this case) for electron microscopy.

“If you fix the cells first, you have a dramatic drop in the efficiency of these immunochemical reactions,” said Igor Kireev, a visiting scientist in the department of cell and developmental biology and lead author of the paper.  Electron microscopy image Click photo to enlarge Image courtesy of Andrew Belmont and Igor Kireev The new technique exposes living cells to labeled antibodies, an approach that yields a much stronger signal for electron microscopy.

“And if your target is inside the condensed chromatin, the antibodies have no way to penetrate.”

Instead of fixing the cells before staining with antibodies, the researchers first exposed living animal cells to the labeled antibodies.  This allowed the antibodies to penetrate more deeply into the chromatin structure, and boosted the number of gold particles adhering to regions of interest.  The signal was enhanced by adding a silver solution that precipitated (solidified) upon contact with the gold.

“We are interested in chromatin structure, so our targets are mostly chromatin-bound proteins,” Kireev said.

The researchers had inserted several copies of a bacterial DNA, called the Lac operator, into the chromosomes.  A bacterial protein, the Lac repressor, recognizes and binds to the Lac operator in living cells.

The researchers combined a Lac repressor protein with another protein that fluoresces green under blue light.  This engineered protein adhered to the chromosomes in regions containing the Lac operator sequences.  Under blue light, these regions fluoresced.  A gold-tagged antibody targeted against green fluorescent protein (GFP) was then microinjected into the nucleus of a living cell, which added a metallic signal that could be boosted with silver.

“All this combined gives us a much better signal, a much stronger signal, with the very best structural preservation,” Kireev said.

The fluorescing protein helped the researchers find the regions of interest in the cells.  These areas were then “immunogold” labeled and targeted for electron microscopy.

In the resulting micrographs the researchers saw enhanced staining of the chromosomes.

“We can now apply this same live-cell labeling method to study at high resolution many different GFP-tagged proteins in the cell cytoplasm or nucleus,” said Andrew Belmont, a professor of cell and developmental biology and senior author of the paper.

“In trying to understand chromosomes, people have largely been limited to low resolution visualization of specific chromosomal proteins using light microscopy,” Belmost said.  “This meant everyone has had to do a lot of guessing of how things are put together, leading in many cases to vague, cartoon models of what are likely to be complicated chromosomal structures carrying out DNA functions such as replication and transcription.”

“Now we hope we can simply look and see the real structure using the more than 10-fold higher resolution of electron microscopy,” Belmont said.  “We are really excited to see what we will find using our new method”

(LA JOLLA, CA) Salk researchers zoom in on genome-wide DNA methylation and transcriptomes at single base resolution. Until quite recently, the chemical marks littering the DNA inside our cells like trees dotting a landscape could only be studied one gene at a time.  But new high-throughput DNA sequencing technology has enabled researchers at the Salk Institute for Biological Studies to map the precise position of these individual DNA modifications throughout the genome of the plant Arabidopsis thaliana, and chart its effect on the activity of any of Arabidopsis’ roughly 26,000 genes.

“For a long time the prevailing view held that individual modifications are not critical,” says Joseph Ecker, Ph.D., a professor in the Plant Biology laboratory and director of the Salk Institute Genomic Analysis Laboratory.  “The genomes of higher eukaryotes are peppered with modifications but unless you can take a detailed look at a large scale there is no way of knowing whether a particular mark is critical or not.”

The Salk study, which appears today in the online issue of Cell, paints a detailed picture of a dynamic and ever-changing, yet highly controlled, epigenome, the layer of genetic control beyond the regulation inherent in the sequence of the genes themselves.

Being able to study the epigenome in great detail and in its entirety will provide researchers with a better understanding of plant productivity and stress resistance, the dynamics of the human genome, stem cells’ capacity to self-renew and how epigenetic factors contribute to the development of tumors and disease.

Discoveries in recent years made it increasingly clear that there is far more to genetics than the sequence of building blocks that make up our genes.  Adding molecules such as methyl groups to the backbone of DNA without altering the letters of the DNA alphabet can change how genes interact with the cell’s transcribing machinery and hand cells an additional tool to fine-tune gene expression.

“The goal of our study was to integrate multiple levels of epigenetic information since we still have a very poor understanding of the genome-wide regulation of methylation and its effect on the transcriptome,” explains postdoctoral researcher and co-first author Ryan Lister, Ph.D.

The transcriptome encompasses all RNA copies or transcripts made from DNA.  The bulk of transcripts consists of messenger RNAs, or mRNAs, that serve as templates for the manufacture of proteins but also includes regulatory small RNAs, or smRNAs.  The latter wield their power over gene expression by literally cutting short the lives of mRNAs or tagging specific sequences in the genome for methylation.

But before Lister could start to unravel the multiple layers of epigenetic regulation that control gene expression, he had to pioneer new technologies that allowed him to look at genome-wide methylation at single-base resolution and to sequence the complete transcriptome within a reasonable timeframe.

Collaborating scientists at the ARC Centre of Excellence in Plant Energy Biology at the University of Western Australia in Perth developed a powerful, web-based genome browser, which played a crucial role in unlocking the information hidden in the massive datasets.

Cells employ a whole army of enzymes that add methyl groups at specific sites, maintain established patterns or remove undesirable methyl groups.  When Lister and his colleagues compared normal cells with cells lacking different combination of enzymes they discovered that cells put a lot of effort in keeping certain areas of the genome methylation-free.

On the flipside, the Salk researchers found that when they knocked out a whole class of methylases, a different type of methylase would step into the breach for the missing ones.  This finding is relevant for a new class of cancer drugs that work by changing the methylation pattern in tumor cells.

“You might succeed in removing one type of methylation but end up with increasing a different type,” says Ecker.  “But very soon we will be able to look and see what kind of compensatory changes are happening and avoid unintended consequences.”

Previous studies had found that a subset of smRNAs could direct methylation enzymes to the region of genomic DNA to which they aligned.  Overlaying genome-wide methylome and smRNA datasets confirmed increased methylation precisely within the stretch of DNA that matched the sequence of the smRNA.  Conversely, heavily methylated smRNA loci tended to spawn more smRNAs.

“We looked at a plant genome but our method can be applied to any system, including humans,” says Lister.  Although the human genome is about 20 times bigger than the genome of Arabidopsis – plant biologists’ favorite model system not least because of its compact genome – Ecker predicts that within a year or so, sequencing technology will have advanced far enough to put the 3 billion base pairs of the human genome and their methyl buddies within reach.

“This really is just the beginning of unmasking the role of these powerful epigenetic regulatory mechanisms in eukaryotes,” says Ecker.