Structure Organization And Function Of The Human Body Biology Essay

Biology » Structure Organization And Function Of The Human Body Biology Essay

Cell are the structural and functional units of all living organisms. Some organisms, such as bacteria, are unicellular, consisting of a single cell. Other organisms, such as humans, are multicellular, or have many cells—an estimated 100,000,000,000,000 cells! Each cell is an amazing world unto itself: it can take in nutrients, convert these nutrients into energy, carry out specialized functions, and reproduce as necessary. Even more amazing is that each cell stores its own set of instructions for carrying out each of these activities.

Prokaryotic Cells - organisms that are lack of nuclear membrane, the membrane that surrounds the nucleus of a cell. Bacteria are the best known and most studied form of prokaryotic organisms, although the recent discovery of a second group of prokaryotes, called archaea, has provided evidence of a third cellular domain of life and new insights into the origin of life itself.

- prokaryotes are unicellular organisms that do not develop or differentiate into multicellular forms.

- are capable of inhabiting almost every place on the earth, from the deep ocean, to the edges of hot springs, to just about every surface of our bodies.

Prokaryotes are distinguished from eukaryotes on the basis of nuclear organization, specifically their lack of a nuclear membrane. Prokaryotes also lack any of the intracellular organelles and structures that are characteristic of eukaryotic cells. Most of the functions of organelles, such as mitochondria, chloroplasts, and the Golgi apparatus, are taken over by the prokaryotic plasma membrane. Prokaryotic cells have three architectural regions: appendages called flagella and pili—proteins attached to the cell surface; a cell envelope consisting of a capsule, a cell wall, and a plasma membrane; and a cytoplasmic region that contains the cell genome (DNA) and ribosomes and various sorts of inclusions.

Eukaryotes include fungi, animals, and plants as well as some unicellular organisms. Eukaryotic cells are about 10 times the size of a prokaryote and can be as much as 1000 times greater in volume. The major and extremely significant difference between prokaryotes and eukaryotes is that eukaryotic cells contain membrane-bound compartments in which specific metabolic activities take place. Most important among these is the presence of a nucleus, a membrane-delineated compartment that houses the eukaryotic cell’s DNA. It is this nucleus that gives the eukaryote—literally, true nucleus—its name.

The outer lining of a eukaryotic cell is called the plasma membrane. This membrane serves to separate and protect a cell from its surrounding environment and is made mostly from a double layer of proteins and lipids, fat-like molecules. Embedded within this membrane are a variety of other molecules that act as channels and pumps, moving different molecules into and out of the cell. A form of plasma membrane is also found in prokaryotes, but in this organism it is usually referred to as the cell membrane.

The cytoskeleton is an important, complex, and dynamic cell component. It acts to organize and maintain the cell's shape; anchors organelles in place; helps during endocytosis, the uptake of external materials by a cell; and moves parts of the cell in processes of growth and motility. There are a great number of proteins associated with the cytoskeleton, each controlling a cell’s structure by directing, bundling, and aligning filaments.

Inside the cell there is a large fluid-filled space called the cytoplasm, sometimes called the cytosol. In prokaryotes, this space is relatively free of compartments. In eukaryotes, the cytosol is the "soup" within which all of the cell's organelles reside. It is also the home of the cytoskeleton. The cytosol contains dissolved nutrients, helps break down waste products, and moves material around the cell through a process called cytoplasmic streaming. The nucleus often flows with the cytoplasm changing its shape as it moves. The cytoplasm also contains many salts and is an excellent conductor of electricity, creating the perfect environment for the mechanics of the cell. The function of the cytoplasm, and the organelles which reside in it, are critical for a cell's survival.

Two different kinds of genetic material exist: deoxyribonucleic acid (DNA) and ribonucleic acid (RNA). Most organisms are made of DNA, but a few viruses have RNA as their genetic material. The biological information contained in an organism is encoded in its DNA or RNA sequence.

Prokaryotic genetic material is organized in a simple circular structure that rests in the cytoplasm. Eukaryotic genetic material is more complex and is divided into discrete units called genes. Human genetic material is made up of two distinct components: the nuclear genome and the mitochondrial genome. The nuclear genome is divided into 24 linear DNA molecules, each contained in a different chromosome. The mitochondrial genome is a circular DNA molecule separate from the nuclear DNA. Although the mitochondrial genome is very small, it codes for some very important proteins.

The human body contains many different organs, such as the heart, lung, and kidney, with each organ performing a different function. Cells also have a set of "little organs", called organelles, that are adapted and/or specialized for carrying out one or more vital functions. Organelles are found only in eukaryotes and are always surrounded by a protective membrane. It is important to know some basic facts about the following organelles.

The nucleus is the most conspicuous organelle found in a eukaryotic cell. It houses the cell's chromosomes and is the place where almost all DNA replication and RNA synthesis occur. The nucleus is spheroid in shape and separated from the cytoplasm by a membrane called the nuclear envelope. The nuclear envelope isolates and protects a cell's DNA from various molecules that could accidentally damage its structure or interfere with its processing. During processing, DNA is transcribed, or synthesized, into a special RNA, called mRNA. This mRNA is then transported out of the nucleus, where it is translated into a specific protein molecule. In prokaryotes, DNA processing takes place in the cytoplasm.

Ribosomes are found in both prokaryotes and eukaryotes. The ribosome is a large complex composed of many molecules, including RNAs and proteins, and is responsible for processing the genetic instructions carried by an mRNA. The process of converting an mRNA's genetic code into the exact sequence of amino acids that make up a protein is called translation. Protein synthesis is extremely important to all cells, and therefore a large number of ribosomes—sometimes hundreds or even thousands—can be found throughout a cell.

Ribosomes float freely in the cytoplasm or sometimes bind to another organelle called the endoplasmic reticulum. Ribosomes are composed of one large and one small subunit, each having a different function during protein synthesis.

2. Describe and distinguish between the cell and tissue organizations and systems.

Tissues are the collection of similar cells that group together to perform a specialized function. The four primary tissue types in the human body: epithelial tissue, connective tissue, muscle tissue and nerve tissue.

Epithelial Tissue - The cells are pack tightly together and form continuous sheets that serve as linings in different parts of the body.  It serves as membranes lining organs and helping to keep the body's organs separate, in place and protected.  Some examples of epithelial tissue are the outer layer of the skin, the inside of the mouth and stomach, and the tissue surrounding the body's organs.

Connective Tissue - There are many types of connective tissue in the body.  It adds support and structure to the body.  Most types of connective tissue contain fibrous strands of the protein collagen that add strength to connective tissue.  Some examples of connective tissue include the inner layers of skin, tendons, ligaments, cartilage, bone and fat tissue.  In addition to these more recognizable forms of connective tissue, blood is also considered a form of connective tissue.

Muscle Tissue - Muscle tissue is a specialized tissue that can contract.  Muscle tissue contains the specialized proteins actin and myosin that slide past one another and allow movement.  Examples of muscle tissue are contained in the muscles throughout your body.

Nerve Tissue - Nerve tissue contains two types of cells: neurons and glial cells.  Nerve tissue has the ability to generate and conduct electrical signals in the body.  These electrical messages are managed by nerve tissue in the brain and transmitted down the spinal cord to the body.

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Subcloning Experiments And Procedures Biology Essay

 

Subcloning is a technique used to produce recombinant DNA. A fragment of DNA containing a gene of interest is inserted into a vector/ plasmid DNA that replicates independently of chromosomal DNA to produce recombined DNA. In order for DNA to recombine, isolation, purification, quantification, digestion, electrophoresis, ligation, transformation, and screening must for all intents and purposes be performed. First alkaline lysis is used to isolate the vector and insert of choice, from bacterial cultures, by opening up the bacterial cell wall and releasing plasmid DNA. Purification removes RNA and protein that may contaminate insert and vector DNA. Quantification reveals the amount of DNA that was isolated (Schramm, Molecular & Cellular Biology Laboratory.). Digesting the DNA sequences with restriction enzymes allows for the extraction of insert and vector DNA at proper sites, which will determine the purity of the DNA samples by using agarose gel electrophoresis (Schramm, Molecular & Cellular Biology Laboratory.). Ligation of the insert to the vector is performed and then transformed into competent cells and grown on plates containing a selectable marker. To determine if the preceding procedures mentioned were successful, the DNA is isolated by doing a restriction digest and the recombinant gene is screened (Schramm, Molecular & Cellular Biology Laboratory). In this particular experiment, if the procedures are carried out successfully the vector will take in the insert gene containing ampicillin resistance and as a result bacterial growth should be seen in the presence of ampicillin and isolation of this recombinant DNA should be possible.

Isolation and purification of DNA from a bacterial culture of E. coli with a vector/plasmid DNA sequence from another bacterial culture of E. coli containing ampicillin resistance, as an insert. An alkaline lysis, with SDS detergent, was used to isolate the DNA from the E. coli cells along with COLD lysis solution. The lysate was incubated at room temperature for 3 minutes and then poured into a spin column and centrifuged, then washed with wash buffer. The DNA was eluted with water, and then underwent centrifugation twice while decanting the flow through, which was then used for quantification while the spin column was discarded. Two vectors and two inserts were used to increase the chances of obtaining purified vector and insert. For quantification, 5 tubes were used, 1 blank (water), 2 separate tubes each containing inserts, and 2 separate tubes containing vector. The blank contained 1000ul of water, each of the 4 tubes contained 5ul of vector DNA and insert DNA and 995ul of water. A spectrophotometer was used to determine the concentration of DNA, and the absorbance. Calculation of the volume of vector and insert was done to obtain 5ug of DNA to be digested by restriction endonucleases Xba1 and BamHI.

Subsequently, 1% agarose gel was prepared with dissolved TAE buffer and ethidium bromide and then solidified. The vector and insert DNA was loaded into the gel. Electrophoresis was done by running the gel for an hour at 120V. Vector1 and insert1 were excised out of the gel as they were highest in concentration of DNA, and then weighed. Three volumes of binding buffer was added to every volume of gel slice and incubated at 50oC for 15 minutes. The original volume of vector and insert of isopropanol was added and mixed by inversion to the DNA samples. Wash buffer was put in and then centrifuged and the filtrate were removed. Centrifugation, elution with 20ul of water, and another centrifugation were done. Next, ligation of the insert to the vector was performed by joining 100ng of vector with no insert (a1:0 ratio), with an equal molar concentration of insert (a 1:1 ratio), and then 3 times the concentration of insert with the vector (a 1:3 ratio). The molar ratio of insert to vector was determined to calculate the volume of the vector and inserts. These volumes were adjusted to 10ul of deionized water. Ligation buffer was added and then mixed. T4 DNA ligase was added and then mixed and centrifuged briefly, then incubated at room temperature for 15 minutes. Later, 5ul of the ligation mixture was transferred into a microfuge tube. Competent cells were added to the DNA and gently mixed by pipeting up and down. The mixture was incubated on ice for 30 minutes and heat shocked for 2 minutes at 37oC, and then cooled on ice for 5 minutes. Luria broth was added, followed by incubation of the cells for 1 hour, 37oC. These mixtures were added to and spread on the appropriate plates. The plates were then incubated at 37oC overnight.

Screening and purification of the recombinant DNA were carried out. Bacterial culture was pelleted, followed by removal of supernatant. This step was repeated with COLD lysis buffer, followed by constant vortexing. The lysate was incubated at room temperature for 3 minutes, and then centrifuged. Wash buffer was added to the tube, followed by centrifugation, removal of the filtrate, decant, and then centrifuged again. The sample was eluted with 30ul of water and centrifuged. Then, DNA quantification was performed. Restriction digest, with the enzyme HindIII, of the quantified recombinant DNA, with a master mix of BSA, Restriction buffer enzyme buffer, and restriction enzyme A, in one microfuge tube was performed. Four more tubes were filled with 3ul of DNA each, and 4ul of H2O, and 3ul of master mix into each of the 4 tubes. These tubes were spun for 1 minute, and incubated at 37oC for 30 minutes. Lastly, 2ul of 5xdyes was added to each tube, so they can be used in electrophoresis, in 1% agarose gel.

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Studying Cells With Super Resolution Microscopies Biology Essay

With a growing interest in biology and the composition of living biological entities as well as a good understanding about the fact that biological entities were composed of extremely small complexes, it was essential to come up with an instrument which would help in viewing objects that could not be seen with an unaided eye. The earliest development of the microscope can be traced back to the use of a magnifying glass even though it wasn’t until the 16th century when the earliest simple microscope was developed by inverting a telescope and this was further modified and improved in the 17th century. Ever since then, new techniques have been developed in order to gain a better understanding of biological entities and presently, the world has reached an era of ‘super-resolution microscopy’ which helps I surpassing ‘Abbe’s resolution limit’. These techniques have helped in imaging nanoscopic molecules that play an essential role in different biological processes and has improved the understanding of the structural and functional properties of subcellular components. Although these techniques have been developed to provide a wide range of properties like 3-dimensional imaging and live imaging, each of them still has its advantages and pitfalls and this essay discussed a few of these techniques in detail.

Introduction

The history of discoveries in cell biology and its related fields is mirrored with the advancements made with the microscope over the past five centuries. Although the simplest microscope was first known to be made and used by Robert Hooke, It was Antonie van Leeuwenhoek who earned the title of “Father of the Microscope” for building the first microscope  in 1674, and pioneering discoveries concerning bacterial cells and erythrocytes. The nineteenth century was marked with improvements in microscopes and staining methods, which further led to scientists establishing the cell theory and viewing the key cell components, understanding cell division and differentiation and the discovery of mitochondria. With the breakthrough of research in the biological field, Ernst Abbe, a mathematician formulated the “Abbe Sine Condition” which enabled calculations that allowed the maximum resolution in microscopes possible  . However, this also meant that that the resolution of optical microscopes was limited by diffraction, which would reach a peak and limit the ability of seeing molecules closely located to one another.

Cell biology revolutionised in the mid-twentieth century with the advances in fluorescent-labelling techniques, which proved to be important tools in biological research, and advances in microscope design and technology. Since then, more specifically in the past decade, there has been an outbreak in the practical implementation of microscopic techniques, with the emergence of super-resolution microscopy that can overcome Abbe’s limit of resolution  , hence converting fluorescence microscopy into an effective 3D visualization tool  . This enables scientists to view single nanoscopic molecules of 10-20nm, not only in all three dimensions, but also trace these molecules in cellular processes. These techniques, as seen in fig.1, follow one of the two approaches; the first is based on spatial patterning of excited light (illumination-based) and this is used in stimulated emission depletion (STED) microscopy and structured illumination microscopy (SIM). The other approach is based on the localization of single molecules (probe-based) and this is used in (fluorescence) photoactivation localization microscopy/Stochastic optical reconstruction microscopy [(f) PALM/STORM]  .

STED

SSIM

TIRFM

PALM/STORM

Best resolution (nm)

~20(lateral)

~50 (spatial)

~50 (lateral)

~90 (spatial)

~230(lateral) ~100 (spatial)

~ 20-30 (lateral)

~60-70 (spatial)

Principle approach

Patterning of excited light using two laser beams

multiple interfering light beams to form moiré patterns

Evanescent field produced by total internal reflection of light

single-molecule localization of photoswitchable fluorophores

Limitations

Photobleaching can occur due to limited light wavelengths

Technical faults

High-intensity pulsed lasers can cause damage to the sample

Photobleaching can occur

Very sensitive to even the smallest changes in sample position

For 3D SSIM, a large number of images need to be taken per wavelength; this takes a long time.

Only 1 plane can be imaged

3D imaging is not possible unless TIRFM is combines with another technique  

Require large number of raw images  

Data acquisition speed is very slow, bearing a direct effect on imaging live samples

Live imaging

TIRFM-SSIM

3D imaging

TIRFM-SSIM

Multicolour imaging

Table 1: Comparison of the distinct features of the different super-resolution techniques.

Two different approaches to breaking the diffraction limit. A. STED microscopy uses two different lasers- an excitation laser (left) and a doughnut-shaped STED laser (middle- this laser deactivates the fluorophores molecules). Using these 2 lasers, the effective excitation area is limited to a small central zone (right). B. Single molecule localization microscopy methods such as PALM and STORM use photoactivatable fluorophores which can switch between their excited state and ground state to successively image the localization of a small number of molecules at a time at high precision by finding the molecule’s centroid. The many ‘raw’ images are then reconstructed to generate the final super-resolution image.

The emergence of super-resolution microscopy has opened many doors in the field of modern biology and medicine, giving an insight on processes that were unable to be followed using conventional microscopy. It is important to understand that every protein found in living cells has a specific function and is a part of a much larger molecular network. In order to understand the functioning of these large networks, it is important to track the movement and interactions of these proteins within the cell  . Super-resolution microscopy, aids in visualising the 3D-structure and accurate location of single protein molecules  on distinct organelles and on structures like lysosomes and microtubules, helping in understanding protein interactions and providing a better understanding of the molecular-scale architecture of cells  .

Three dimensional STORM image of the mitochondria network in a mammalian BS-C-1 cell. The z-position is colour-coded according to the colour scale bar.

In the past decade, super-resolution microscopy has been used to map the 3D-organization of distinct components of the nuclear pore complex; the polygonal network that makes up the endoplasmic reticulum in cells was imaged, as seen in fig.3, in living PtK2-cells of the kidney; the movement of synaptic vesicles have been traced inside living neurons by tagging the vesicle protein synaptotagmin with antibodies  and these techniques have been used to study the co-localization of two mitochondrial proteins by labelling them with different fluorophores. These studies would not have been possible without nanoscale-resolution provided by these techniques since all these structures are extremely small in size  .

Super-resolution imaging of the endoplasmic reticulum in living PtK2-cells of the kidney cell. (A) Shows the confocal image and (B) shows the simultaneously recorded STED (x, y) images from the ER marked by the fluorescent protein Citrine targeted to the ER. The arrows point out rings formed by the tubular network of the ER, which are clearly visible only in the STED image (B). 

The emergence of super-resolution microscopy has put light on important details of cell biology, holding great importance for research in the future. This essay discussed the different techniques of super-resolution microscopy, its application in cell biology, and its limitations as an instrument.

The first super-resolution microscopy technique, STED microscopy’s concept was introduced a decade ago and has advanced within the past few years. It is based on patterning the excited light in such a way that the volume of light in the excited-state is extremely small, hence maintaining the amount of light that emits fluorescence to small volumes  .

This is achieved by using two pulsed laser beams of different wavelengths; the wavelength of light from the first laser beam excites the fluorescent marker and the second laser beam illuminates the sample with a doughnut-shaped beam (called the STED-beam)  as seen in fig.4. The wavelength of light from the STED-beam is such that it causes the excited fluorescent molecules to de-excite, bringing them back to the ground-state via stimulated emission. The doughnut-shaped beam from the second laser ensures that the molecules of the centre-most part of the labelled sample are in the excited state, and fluorescence is detectable  .

Schematic diagram showing the use of the excitation and deexcitation (STED) beams for 3D-STED imaging inside a living cell. (A) An objective lens focuses the excitation (blue) beam and deexcitation (orange) beam into the ER while also collecting the resulting beam from the fluorescence photons. (B) xy-axes imaging: excitation spot (blue) and doughnut-shaped focal spot (orange) for stimulated emission (C) xz-axes imaging: excitation spot (blue) and STED spot composition consisting of a spot featuring a maximum above and below the focal plane along the z- axis, referred to as STEDz, and an enlarged doughnut-shaped beam called STEDr.

The lateral resolution of STED microscopy has been pushed to below 20nm and has been successful in imaging the synaptic vesicle movement in live neurons after neurotransmitter release during an impulse. In the past, synaptic vesicle exocytosis was suggested and confirmed by using electron microscopy, where ‘pockets’ in the pre-synaptic membrane terminals of chemically-fixed nerve cells were seen, hinting on exocytosis as the process of neurotransmitter release  . Further, fluorescence microscopy was used to study the vesicular movement after neurotransmitter release by using FM-dyes  . Even though it was known that vesicles are recycled via endocytosis, the fate of its components after fusion with the membrane was still unclear since the vesicles were too small to be resolved by available microscopes.

To solve this problem, STED microscopy was used to determine the entire process of vesicle endocytosis. Monoclonal antibodies against the intravesicular membrane protein synaptotagmin was used for imaging purposes; these antibodies only bound to those protein molecules that were exposed during vesicle exocytosis and were internalised when the vesicle was endocytosed. Fluorescent-labelled secondary antibodies were attached after membrane fixation and permeabilization and were used for visualisation of these vesicles. Images showed synaptotagmin molecules clustered on the pre-synaptic membrane, suggesting that vesicle components remain together on the pre-synaptic membrane during recycling by endocytosis. Each synaptic vesicle is 40-50nm in size and they usually occur in groups of 100-300 vesicles. Therefore, fig.5 shows that using STED microscopy was essential for localising individual vesicles, and contrary to previous beliefs that vesicles hardly move, STED revolutionised the understanding of vesicle-recycling by showing that vesicles constantly move rapidly and randomly  .

Comparison of confocal (left) and STED (right) counterpart images of a small region of a neuron terminal labelled with an anti-synaptotagmin antibody, ?xed, permeabilized and visualized using Atto532-labelled secondary antibodies. The STED image reveals a marked increase in resolution and also shows the accurate location of individual vesicle components on the neuron membrane.

However, STED microscopy is limited by wavelength. The absence of sufficient tuneable pulsed light sources in the visible range of light which de-excite the already excited fluorescent-labelled molecules has limited STED microscopy to only a small fraction of fluorophores, which causes bleaching and phototoxicity  . STED also requires the use of high intensity pulsed lasers which can cause significant damage to the samples. Furthermore, there are technical limitations set by the laser power required  for this technique and the very often, mechanical drift of the optical instruments causes imperfection of the doughnut-shaped beam around the sample, limiting the spatial resolution.

Another example of an illumination-based technique, SSIM follows the approach of illuminating the sample with multiple interfering light beams in order to break the resolution barrier  . When multiple beams of mutually coherent light are allowed to interfere, they form a structured pattern, like that of Moiré fringes seen in fig.6. When focussed on the labelled sample, the illumination pattern further interacts with the fluorescent probes. The emitted light contains image details of higher resolution, including details that cannot be resolved using a normal microscope. The illumination patterns are modulated by changing the orientation of light on the sample and high-resolution images are captured within the illumination from different patterns.

The approach of resolution enhancement followed by structured illumination. (a) If two line patterns are superposed in each other, moiré fringes will be formed as a product (seen here as the apparent vertical stripes in the overlap region). (b) A conventional microscope is limited by diffraction to a circular ‘observable region’ of reciprocal space. (c) A sinusoidally striped illumination pattern-the possible positions of the two side components (light beams) are limited by the same circle that defines the observable region (dashed). If the sample is illuminated with structured light, moiré fringes which represent information that has changed position in reciprocal space will appear. The observable region will contain normal information and moved information that originates in two offset regions (d). From a series of images with different orientation and pattern phase, it is possible to recover information from a region that is twice the size of the normally observable region can be obtained, corresponding to twice the normal resolution (e).

The images are collected and reconstructed using computer software which extracts the details from the moiré images, reconstructing them into 3-dimensional images with doubled resolution. The original 2D-SIM involved using two beams of light which interacted with the sample probe to increase its resolution and form 2D images. However, this technology was extended by using 3 light beams, generating resolved images with finer details of the sample in the axial and lateral directions, resulting in a 3-dimensional image of the sample.

Using 3D-SIM in comparison with conventional wide-field epifluorescence-microscopy, experiments to better the understanding of higher order chromatin and to study the accurate localizations of other nuclear components like the nuclear pore complexes (NPCs) and nuclear lamina were performed. The chromatin of formaldehyde-preserved mouse C2C12-myoblast cells were stained with 4',6-diamidino-2-phenylindole (DAPI) and they were observed using 3D-SSIM. The Images obtained from this technique showed a large number of ‘holes’ within the area of the stained chromatin as in fig.7, a feature that could not be observed in the images obtained by wide-field epifluorescence-microscopy.

Comparison of 3D-images obtained from conventional wide-field microscopy (left) and 3D-SIM (right) used in order to resolve interphase chromatin structure of the same DAPI-stained C2C12 cell nucleus. Deconvolution was applied to the wide-field data set (middle). (A) Mid cross-section shows brightly stained clusters of centromeric heterochromatin. Inset shows higher-detail information of chromatin substructures when recorded with 3D-SIM. Arrow in 3D-SIM inset points to a less-bright chromatin structure that has been spuriously eroded by the deconvolution procedure. (B) Apical sections (corresponding to a thickness of 0.5 µm) taken from the surface of the nuclear envelope closest to the coverslip. The raw image shows diffuse DAPI-staining, the deconvolved image shows more pronounced variations in fluorescence intensities and the image obtained from 3D-SIM shows extended resolution and reveals a punctuated pattern of regions without DAPI-staining.

Further taking advantage of SSIM’s multicolour and 3D-imaging properties, the same cells were immunostained with antibodies specific to the nuclear pore complexes (NPC), which detect the NPC proteins; and antibodies against lamin-B, a major component of the nuclear lamina (intermediate filament protein). Hence, the 3D-SSIM images showed the chromatin on the nucleoplasmic side, followed by nuclear lamina and then the nuclear pore complexes on the cytoplasmic side forming a triple-layered organization as in fig.8. Not only was the heterochromatin distinguished from the euchromatin, but at every ‘hole’ where DAPI-labelled chromatin was absent, some amount of NPC-staining was present, suggesting that chromatin was absent within close proximity of the NPCs. Even though all three sub-nuclear structures were obtained using conventional fluorescence microscopy, the spatial organization of these structures was obtained only by using 3D-SSIM.

multicolour imaging of DNA, nuclear lamina, and NPC structures in C2C12 cells by 3D-SIM. The cells are immunostained with antibodies against lamin B (green) and antibodies that recognize different NPC epitopes (red). DNA is counterstained with DAPI (blue). The image on the top left shows the same sample imaged using confocal laser scanning microscopy (CLSM) and the image on the top right shows the images obtained using 3D-SSIM which are better resolved and more clearer. The bottom picture clearly shows the triple layered organization of the three structures.

Therefore, 3D-SSIM has proved to be essential in understanding the spatial organization and interactions of sub-cellular structures that were unable to be studied before. Even though some of the initial limitations of SSIM like the time required to reconstruct and analyse the images have been overcome, SSIM is still restricted by the photostability of the fluorophores used since photobleaching leads to a less intensive image.

Even though total internal reflection fluorescent microscopy (TIRFM) was first used in 1981, it’s still a very important technique and has been used extensively since it allows selective excitation of labelled molecules in a cellular/aqueous environment which are near the surface only. This is not only beneficial because of its ability to view labelled molecules, but also because the region of interest is thin enough to obtain the highest frame-rates. TIRFM combined with structured illumination microscopy (SIM) has developed into a super-resolution technique which can break the resolution barrier and improve resolution of the region of interest.

The conventional-TIRFM is based on the diffraction properties of a light beam when incident onto a surface separating two media with different (high and low) refractive indexes. At a high incident angle (greater that the critical-angle), all the incident light is ‘totally reflected’ as long as it is coming from the medium with a high refractive index through the medium with a low refractive index  . At this surface, an ‘evanescent field’ is produced. This field is considered to be an electromagnetic field capable of exciting fluorophore molecules present on the surface. This field rises from the surface into the medium of lower refractive index  . The depth of fluorophore excitation is minimised in this phenomenon because as the evanescent field rises parallel to the surface and the distance between the field and surface increases, its strength decreases exponentially, limiting the fluorescent region. TIRFM not only provides a very thin, sectioned layer of excited fluorophores which helps in minimising the background noise caused by water molecules, it also omits unwanted fluorescence of molecules that are out of focus. However, the major drawback of this technique is that only one plane (z-plane) can be imaged, limiting its use to study cell surface events. Therefore, to obtain limit-breaking resolutions and to view multiple planes of the sample region, TIRFM is used in combination with SIM.

The TIRFM-based SIM was used to image EGFP-labelled a-tubulin of living S2-cells of Drosophila. a-tubulin is a protein present in microtubules. Comparing the images of the same sample region obtained by using conventional-TIRFM and TIRFM-SIM, the latter showed a significant improvement in the resolution of the image as seen in fig.9(a,b).

Comparison of conventional TIRF (a) and TIRF-SIM (b) images of the microtubule cytoskeleton in a single S2 cell. The image obtained after combining TIRF and SIM shows better resolution hence giving a clearer image.

Live imaging using TIRFM-SIM was applied to image polymerisation-depolymerisation of microtubules located near the centrosome of a Drosophila S2-cell which was in its mitotic state. Since the length of microtubules was constantly changing due to its polymerisation and depolymerisation, kymographs were used to process images and to determine the spatial-position of the microtubules over time by determining the difference in GFP-labelling density along the microtubule length at different times. Combing SIM with TIRFM helped in imaging the GFP-labelled a-tubulin with enhanced clarity and allowed accurate localization of the end of the microtubule, hence being able to follow it through the process. The images obtained from live-SSIM showed distinct transformation between the microtubule’s polymerisation state, depolymerisation state and its steady state, hence being able to track the dynamics of the microtubules (fig.10), a phenomenon which was not possible to understand properly using conventional-TIRFM.

TIRF-SIM images at different time frames of EGFP-a-tubulin in a S2 cell. (a) 95th image from a 180-frame sequence. Each frame was acquired in 270ms. (b) The green-boxed area of (a) shown at selected times as indicated on the individual images, using conventional TIRF (left) and TIRF-SIM (right). Green arrows follow the end of a single microtubule, which can be seen elongating until approximately the 100 s time point, and then rapidly shrinking. These changes are much easier to follow in the TIRFM-SIM images which are much clearer compared to the TIRFM images obtained.

In contrast to STED and SIM-microscopy (based on the spatial patterning of excited light), STORM and photoactivated localization microscopy (PALM) are probe-based methods principled on single-molecule localization and were developed recently in 2006. These techniques combine 3D and multicolour-imaging and obtain images with a spatial-resolution of 20-30nm and an axial and lateral-resolution of 60nm and 70nm respectively  . Keeping in mind that single molecule localization is made difficult in fluorescently-labelled biological samples because it contains millions of fluorophore molecules in a large density  , PALM/STORM use photoswitchable probes which can be switched between its visible (fluorescent, excited) and invisible (nonfluorescent, de-excited) state by using light of different wavelengths. Therefore, this approach consists of repeated cycles of sample imaging. In each cycle, different fluorophore-molecules within a diffraction-limited region are excited, such that each excited molecule can be individually imaged without overlapping (due to the images of closely located fluorophore molecules which are invisible in this cycle) and subsequently deactivated to the ground-state  as seen in fig.11. In following cycles, a stochastically different set of fluorophore-molecules are excited, determining the accurate coordinates of different molecules in each image. Using these individual images, an overall image is constructed and the position of each molecule in the sample is determined. The PALM/STORM techniques and based on the same concept of single-molecule localization, the only difference being the fluorescent probes that each of them uses. While PALM originally used photoactivable fluorescent proteins that are attached to sub-cellular structures, STORM used synthetic photoswitchable cyanin dyes that carried out the same function.

Schematic diagram showing the basic principle followed by STORM imaging. (a) Shows the microtubules within a cell. (b) shows a distinct set of fluorophore molecules in its excited state. (c), (d) and (e) show different set of fluorophore molecules that are excited while the other closely situated molecules are in the ground-state by their photoswitchable property. (f) shows the complete reconstructed image formed by compiling all the raw images into one image.

Further, STORM is developed to provide multi-coloured imaging by using combinatorial pairs of “reporter” dyes which cause the fluorescence and “activator” dyes which can reactivate the ‘switched-off’ reporter dyes when placed in close proximity to the reporter. Thus, each pair has a different colour of emitted light, determined by the reporter dye and a different colour light that activates the reporter, determined by the activator dye  . This technique, therefore, allows the study of molecular interactions between different sub-cellular structures by co-localizing them within a cell.

Comparison between images of microtubules in a mammalian cell obtained from conventional microscopy and 3D-STORM (A) Conventional immunofluorescence image of microtubules in an area of a BS-C-1 cell. (B) The 3D-STORM image of the same area with the z-position of the microtubules colour-coded according to the colored scale bar. (C-E) Show the x-y, x-z and y-z cross-sections of a small region of the BS-C-1cell outlined by the white box in (B), showing 5 microtubule filaments.

To understand the interaction and spatial relation between mitochondria and microtubules within a cell, a two-colour 3D-STORM was performed which proved to be fundamental towards the understanding of mitochondrial-microtubule interactions. It is certain that mitochondria are the “power houses” of a cell and hence, to maintain its dynamic morphology  , these organelles are constantly moving about a cell with help from motor-proteins which attach particularly to microtubules within a cell. For this experiment, fixed monkey kidney BSC-1-cells were used and two different sets of reporter-activator dyes were used to stain Tom20, part of the translocase outer mitochondrial membrane complex (used as an outer membrane marker for mitochondria) and ß-tubulin, a protein present in microtubules; the reporter dyes were attached to secondary antibodies.

Comparison of images of microtubule-mitochondrial interactions in mammalian cells as obtained from conventional and 3D-STORM microscopy. (a) A conventional fluorescence image of mitochondria (magenta) and microtubules (green). The image is slightly blurred and the distance between the mitochondria and microtubules, if any, is not visible since a single mitochondrion is seen to touch multiple microtubules. (b) STORM image of the same area with all localizations at different z positions stacked. The image is acquired in aqueous media and reconstructed from 500,000 localization points. This image, contrary to the conventional image (a), clearly shows a 150nm separation between the mitochondrion and one microtubule, whereas the same mitochondrion was in much closer proximity to another microtubule.

The STORM-image provided a clear picture of mitochondria and microtubules, allowing a better understanding of the spatial relation between them as compared in fig.13. STORM is not limited to imaging the interactions of only two sub-cellular structures and can be used to image multiple structures by differentially labelling them. STORM can be extended to imaging motor-proteins, the main complexes which facilitate the mitochondrial movement along microtubules, further illustrating their interactions and providing a better understanding of the regulation of morphology of these “power houses” within a cell, thereby having much potential for future nanoscale-research.

However, STORM requires large numbers of raw images of localised-molecules to be taken from different imaging-frames so that the entire super-resolution image can be constructed, and this limits the speed of this technique and the acquisition time required to construct the highest-resolution image requires a few minutes  .

Conclusion

It is safe to conclude by saying that microscopy has come a long way since its first discovery in the late 16th century and reached an era when the diffraction limit is being surpassed so that individual nanoscale molecules can be observed. In the past decade, super-resolution microscopy has taken a big jump and techniques like STED, SSIM, TIRF-SSIM and PALM/STORM have been developed. Even though each of these techniques accommodates features like greatly improved image-resolution, 3-dimensional imaging, live-sample imaging and multi-coloured imaging, each of these has its own limitations. In the ideal world, STED microscopy would be expected to work independent of light-wavelength and unaffected by the high-intensity lasers. Similarly, PALM/STORM would be expected to be faster techniques requiring lesser raw-images and SSIM would be expected to be unaffected by photobleaching and sample-positioning. SIM can be applied for live-imaging, 3D-imaging and multicolour-imaging; however, its resolution is still not as good as that provided by STED microscopy (live-imaging, 3D-imaging) and PALM/STORM (3D-imaging, multicolour-imaging). Therefore, at this point, it is hard to tell as to which of the above explained techniques is the best since each of them have their advantages and pitfalls and each has significant potential in different areas of biological research. As of now, considerable progress has been made in microscopy, hence opening many doors in cell biology and it is safe to say that in the future, technology will improve, and new imaging techniques will be developed

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Structure Function Studies Of Helicobacter Pylori Urease Biology Essay

Biology » Structure Function Studies Of Helicobacter Pylori Urease Biology Essay

Urease catalyzes the hydrolysis of urea into ammonia and carbon dioxide. The release ammonia neutralizes the gastric acid, and allows the colonization of Helicobacter pylori in human stomach. The apo-urease undergoes a post-translation carbamylation of an active-site lysine residue, followed by insertion of two nickel ions essential for metal catalysis to the active site. In H. pylori, four urease accessory proteins, UreE, UreF, UreG, and UreH, are essential to the maturation process of urease. It is postulated that the apo-urease either bind a pre-formed UreG/UreF/UreH complex, or the individual urease accessory proteins sequentially to form a pre-activation complex. The Ni-binding protein UreE then interacts with the UreG of the complex, and triggers the GTP-dependent activation of urease.

How these urease accessory proteins interact with each other and with the urease to form the activation complex is poorly understood, partly because of the lack of high-resolution structures available for these urease accessory proteins. Until recently, the only urease accessory protein whose structure is available is UreE. We have recently determined the crystal structure of UreF, and have obtained a preliminary structure of the UreF/UreH complex. The novel structural information allows us to use the protein engineering approach to address the following questions: (1) Is the dimerization of H. pylori UreF/UreH complex essential to the maturation of urease? (2) Is the interaction between UreF and UreH essential to the maturation of urease? (3) Where is the interacting surface on the UreF/UreH complex that are responsible for binding UreG and the urease? As we have already obtained crystals of the UreG/UreF/UreH complex that diffract to good resolution, we propose to determine the crystal structure of this ternary complex. Through this work, the structures of all urease accessory proteins involved in urease maturation will be available, and together with the mutagenesis data, we will have a better understanding of how the urease accessory proteins associate with the urease to form the pre-activation complex for the maturation of urease.

Infection of Helicobacter pylori induces inflammation in the human stomach, and causes gastric or duodenal ulcers. High activity of urease is one of the factors that facilitate the colonization of H. pylori in the stomach. Urease catalyzes the hydrolysis of urea into ammonia, which neutralizes the gastric acid and allows the pathogen to survive in the human stomach.

In the active site of urease, there is a carbamylated lysine residue that are involved in binding two nickel ions essential to the metal catalysis of the enzyme. In H. pylori, the maturation of urease (i.e. the carbamylation of the active site lysine residue and the insertion of nickel ions) is assisted by four urease accessory proteins, namely, UreE, UreF, UreG, and UreH (UreH is the H. pylori ortholog of UreD found in other species). The current model for urease maturation suggests that the urease binds UreF, UreG and UreH to form a pre-activation complex, which then interacts with the Ni-binding protein UreE to trigger the GTP-dependent activation of urease.

How the urease accessory proteins interact with each other and with the urease for the maturation process is poorly understood. This proposal aims to address a number of important questions concerning the structure-function of urease accessory proteins UreF and UreH:

(1) Is the dimerization of H. pylori UreF/UreH complex essential to the maturation of urease?

We have determined the crystal structure of H. pylori UreF and UreF/UreH complex. Both of them exist as dimers in the crystal structure. Moreover, we showed that H. pylori UreF/UreH complex exists as a 2:2 dimer in solution. Based on the crystal structures, we will introduce mutations in the dimeric interface to break the dimerization of H. pylori UreF/UreH complex, and test if these mutations affect the in vivo maturation of urease.

(2) Is the interaction between UreF and UreH essential to the maturation of urease?

Our preliminary data showed that the truncation of the C-terminal residues of UreF breaks the UreF/UreH complex, and abolishes the activation of urease. To further investigate the functional importance of UreF/UreH interaction in urease maturation, we will introduce mutations in the interface of the UreF/UreH complex, and test if these mutations affect the formation of UreF/UreH complex, and in vivo maturation of urease.

(3) How does the UreF/UreH complex interact with UreG and the urease?

The UreF/UreH is also known to form bigger complexes with UreG, and with the urease. How UreF/UreH complex associates with UreG and the urease to form the activation complex for the maturation of urease is poorly understood. Based on the crystal structure of the UreF/UreH complex we solved, we propose to perform scanning mutagenesis to map the surface on UreF and UreH for interaction with UreG and with the urease. The mutants’ ability to form complex with UreG and the urease will be correlated with their ability to activate urease in vivo.

(4) What is the structure of UreG/UreF/UreH complex?

We have already obtained crystals of UreG/UreF/UreH complex. We propose to solve the high-resolution structure of the complex by X-ray crystallography. The structure will provide the first high-resolution structure of how these urease accessory proteins interact with each other.

Through this work, we will determine the crystal structure of UreF/UreH/UreG complex. Together with the mutagenesis data and the in vivo urease activation assay, our proposed work will contribute a significant step towards a better understanding on the structure-function relationship of these urease accessory proteins.

Objectives

To test whether the dimerization of the H. pylori UreF/UreH complex is essential to the maturation of urease

To investigate the functional interaction between UreF and UreH by mutagenesis.

To map the interaction surface on the UreF/UreH complex for binding UreG and the urease by scanning mutagenesis.

To determine the structure of UreG/UreF/UreH complex by X-ray crystallography

Activity of urease is one of the factors that facilitate colonization of Helicobacter pylori in the human stomach. Urease is a nickel-containing enzyme that hydrolyzes urea into ammonia and carbamic acid, which decomposes spontaneously into carbonic acid and ammonia [1].

NH2-CO-NH2 + H2O ? NH3 + NH2-COOH

NH2-COOH + H2O ? NH3 + H2CO3

The ammonia neutralizes the gastric acid, and allows the pathogen to survive in the human stomach.

The structure of urease from various species have been determined [2-5]. Urease is composed of ?, ? and ? subunits. In H. pylori, the ureA gene encodes the b and g subunits as a fusion protein, and the ureB gene encodes the a-subunit. In the active site of urease, a carbamylated lysine residue is involved in binding two nickel ions, which are essential to the catalysis of urea hydrolysis. The maturation of urease involves the carbamylation of the lysine residue and the insertion of nickel ions to the active site. In H. pylori, the urease accessory proteins that are involved in urease maturation are: UreE, UreF, UreG and UreH [1]. UreH is the H. pylori ortholog of UreD found in other species. In this proposal, we use the notation of ‘UreH(D)’ when we refer in general to the homologous UreH or UreD proteins, and use ‘UreH’ when we refer specifically to the protein UreH in H. pylori.

UreF and UreH(D) play pivotal roles in the formation of activation complex for the maturation of urease. UreF was reported to form complex with UreH(D) [6-9], and the two proteins interact with UreG to form the heterotrimeric complex UreG/UreF/UreH(D) [9, 10]. UreG is a SIMIBI class GTPase, which is homologous to the hydrogenase maturation factor HypB [11]. The apo-urease can form complex with UreG/UreF/UreH(D), or its components of UreH(D) and UreF/UreH(D) [9, 10, 12, 13]. It has been shown that apo-urease can be activated in vitro by just adding excess amount of carbon dioxide and nickel ion [14]. The in vitro activation of urease is increased when in complex with UreF/UreH(D) and UreG/UreF/UreH(D) [13, 15]. Chemical cross-linking experiments suggest that binding of UreF/UreH(D) may induce conformational changes of the urease [16], which may allow the diffusion of nickel ion and carbon dioxide into the active site to promote activation of urease [17].

The current model for in vivo urease maturation proposed by Hausinger’s group is illustrated in Fig. 1 [1]. The apo-urease interacts with UreG, UreF and UreH(D) to form a pre-activation complex. UreE, a dimeric nickel-binding protein, then interacts with UreG of the complex, and triggers the GTP-dependent activation of urease [15, 18, 19].

The formation of activation complex for the maturation of urease involves protein-protein interaction among the urease accessory proteins and the urease. However, structure-function studies of how these urease accessory proteins interact with each other was only poorly understood. One obstacle was that expression of UreH(D) alone in E. coli resulted in the formation of inclusion bodies. Recently, Hausinger’s group has successfully expressed soluble K. aerogenes UreD in fusion with the maltose binding protein (MBP-UreD), which allows for the first time in vitro characterization of UreH(D). They showed that UreH(D) can interact with UreF in ~ 1:1 binding ratio, but only weakly with UreG [9].

Until recently, the only urease accessory protein whose structure is available is UreE [18, 20-22]. The structure of UreF was recently determined by Chirgadze’s group [23], and in parallel, by our group [24]. The work proposed here will fill the much-needed gap of knowledge on the structure-function studies of urease accessory proteins.

1. We have determined the crystal structure of H. pylori UreF. Our group has determined the crystal structure of H. pylori UreF using the MAD method with Se-Met labeled protein [24]. The structure of the native UreF, refined to 1.85 Å resolution by us, is similar to the structure of Se-Met derivative reported independently by Lam et al. [23]. UreF is an all-alpha protein consisting of 10 helices. It forms dimers in the crystal structure (Fig. 2). The dimeric interface is formed by docking of helix-1 to the helix-8 and helix-9 of the opposite UreF molecule.

2. We have established an efficient protocol to express and purify UreF/UreH complex. As mentioned above, one obstacle for the structure-function studies of the UreF/UreH complex was that expression of UreH alone resulted in insoluble inclusion bodies (Fig. 3). We have successfully solved this problem by co-expressing UreH with GST-UreF in E. coli (Fig. 3). After affinity chromatography purification and removal of the GST-fusion tag, the UreF/UreH complex can be purified in large quantity (~10 mg per liter of bacterial culture).

3. We have established assays to correlate in vitro protein-protein interactions with in vivo maturation of urease. We showed that when co-expressed together, UreF and UreH form a soluble complex that can be pull-down by GST affinity column (Fig. 4A, lane 2). We noticed that the C-terminal residues of UreF were protected from degradation upon complex formation with UreH (Fig. 4A, lane 1 & 2). We showed that truncation of the C-terminal residues of UreF (UreF-DC20) disrupted the formation of a soluble UreF/UreH complex (Fig. 4A, lane 3). We have also established an assay to test the in vivo maturation of urease (Fig. 5), and showed that the mutation (UreF-DC20) that disrupted the interaction between UreF and UreH also abolished in vivo maturation of urease. By GST pull-down, we demonstrated that the UreF/UreH complex interacts with UreG (Fig. 4B, lane 4), and with the urease (Fig. 4B, lane 4). These preliminary data demonstrated that feasibility of the proposed structure-function studies.

4. We have obtained the preliminary crystal structure of H. pylori UreF/UreH complex. With the purified UreF/UreH complex, we were lucky to obtain crystals of the complex that diffract to high resolution (Fig. 6A). Diffraction data was collected to 2.5Å resolution. We phased the structure by molecular replacement using the structure of UreF as a search template. Our preliminary structure of UreF/UreH complex showed that the UreF/UreH complex forms a 2:2 dimer in the crystal structure (Fig. 6B). We anticipate that the refinement of the UreF/UreH complex structure will be finished very shortly, and the structure will provide a rational based for the mutagenesis studies proposed in this study.

5. We have showed that the UreF/UreH complex form dimers in solution. To test if the UreF and UreF/UreH form dimers in solution, we have loaded purified samples of UreF and UreF/UreH to an analytical size-exclusion-chromatography column coupled to a static light scattering detector (Fig. 7). The apparent M.W. for UreF was 43 kDa, which is in between the theoretical M.W. of a monomeric (28 kDa) and a dimeric (56 kDa) form of UreF. The results suggest that UreF alone does have a tendency to form dimers, and the dimeric form of UreF is in exchange with the monomeric form in solution. On the other hand, the formation of dimer is more-or-less complete in the UreF/UreH complex. The apparent M.W. measured for UreF/UreH complex was 116 kDa, which is consistent with the theoretical M.W. of 116 kDa for a 2:2 dimer of UreF/UreH complex.

6. We have established an efficient protocol to express and purify UreG/UreF/UreH complex, and obtained crystals of the complex. We have found that the most efficient way to obtain the H. pylori UreG/UreF/UreH complex is to co-express UreG, GST-UreF and UreH together in E. coli. The ternary complex can be easily purified by affinity chromatography followed by removal of GST-fusion tag by protease digestion. In our hand, the yield of UreG/UreF/UreH complex is ~5mg per liter of bacterial culture. More encouraging is that we have successfully obtained crystals of UreG/UreF/UreH that diffracted to a reasonable resolution of ~3Å (Fig. 8). These preliminary data strongly suggest that the proposed structure determination of the ternary complex of UreG/UreF/UreH by X-ray crystallography is highly feasible.

Track Record of PI

The PI has extensive experience on structure determination by both NMR and X-ray crystallography, and using protein engineering to probe the structure-function of proteins. In addition to the structure determination of UreF and UreF/UreH complex discussed above, he has solved the solution structure of barstar, an inhibitor of barnase, and studied its dynamics behavior by NMR spectroscopy [25, 26]. He also studied the effect of mutations on the stability and structural perturbation on the DNA-binding domain of the tumor suppressor p53 by NMR spectroscopy [27, 28]. He has used an approach that combines evidence from NMR experiments and molecular dynamics simulation to study the folding pathway and the denatured states of barnase and chymotrypsin inhibitor-2 [29-31]. Supported by previous GRF grants, he solved the solution [32] and crystal [33] structure of ribosomal protein L30e from Thermococcus celer, the crystal structures of a thermophilic acylphosphatase from Pyrococcus horikoshii to 1.5Å [34], and human acylphosphatase to 1.45Å [35], an orange fluorescent protein from Cnidaria tube anemone to 2.0Å [36], seabream antiquitin to 2.8Å [37], the crystal structure of trichosanthin in complex with the C-terminal residues of ribosomal stalk protein P2 to 2.2Å [38], and the solution structure of the N-terminal dimerization of P2 [39]. We believe that, with our strong background in structural biology and the solid preliminary data, we are in a leading position to determine the structure of the UreG/UreF/UreH ternary complex, and to study the how the urease accessory proteins interact with each other for the maturation of urease.

Our preliminary data suggest that the H. pylori UreF/UreH complex forms a 2:2 dimer in solution. Both the crystal structure of H. pylori UreF, and the preliminary structure of UreF/UreH complex suggest that the dimerization is likely to be mediated by UreF. It is presently not known whether the dimerization is a unique property of H. pylori UreF - for example H. pylori and K. aerogenes UreF only share 19% sequence identity. Interestingly, the quaternary structure of H. pylori urease is different from ureases from other bacterial species. Unlike the urease (UreABC) from K. aerogenes that forms a trimeric complex (UreABC)3, the H. pylori urease (UreAB) forms a tetramer of trimers ((UreAB)3)4. Nevertheless, that H. pylori UreF/UreH complex exists as a dimer in solution and in crystal structure raises an interesting question - is the dimerization of H. pylori UreF/UreH complex essential to the maturation of urease?

To address this question, we will introduce mutations that are designed to break the dimerization of UreF and UreF/UreH complex. As shown in Fig. 2, the dimeric interface is formed by docking of helix-1 to the helix-8 and helix-9 of the opposite UreF molecule. A closer look at the dimeric interface of the crystal structure of UreF reveals a number of interactions that may be importance to the dimerization of UreF (Fig. 9). For example, to break the hydrogen bonding network among Q37, Q205 and Q212, we will replace the Gln residue with either alanine or asparagine to create triple mutants of Q37A/Q205A/Q212A and Q37N/Q205N/Q212N. We anticipate that both truncation of and shortening of the amide chain should break the hydrogen bond network. To disrupt the hydrophobic interaction around F33, we will substitute the Phe residue with alanine (F33A) or with a polar residue (e.g. F33R). Substitution of polar residue like arginine at Phe-33 should highly disfavor dimerization because the high desolvation penalty will prevent the polar residue to be buried upon dimerization. If necessary, we will create quadruple mutants (e.g. Q37A/Q205A/Q212A/F33A) to ensure disruption of UreF dimerization.

3.1.1 GST pull-down assay for UreF/UreH interaction. First, we test if these mutants will affect the formation of soluble UreF/UreH complex by GST pull-down assay (Fig. 4A). UreH will be co-expressed with mutants of UreF fused with GST-tag, and the bacterial lysate will be loaded to a GSTrap column (GE Healthcare). After extensive washing with binding buffer (20 mM Tris pH7.5, 0.2M NaCl, 5mM DTT), the proteins will be eluted with 10mM glutathione.

As these mutations are located at the dimerization interface, which are far away from the UreF/UreH interface, we anticipate that they will not affect UreF/UreH interaction.

3.1.2 Size-exclusion-chromatography/static-light-scattering (SEC/LS). We will test if these mutants affect dimerization of UreF by SEC/LS. Purified samples of UreF mutants and its complex with UreH complex will be loaded to an analytical Superdex 200 column connected to an online miniDawn light scattering detector and an Optilab DSP refractometer (Wyatt Technologies). The light scattering data will be analyzed using the ASTRA software provided by the manufacturer to obtain the molecular weight of the protein samples.

If the mutations break the dimerization, we anticipate that the measured molecular weight will be 28 kDa for UreF, and 58 kDa for UreF/UreH complex.

3.1.3 In vivo maturation of urease. We will test if test if these mutants affect in vivo maturation of urease. We have established an assay for in vivo maturation of urease (Fig. 5). We have cloned the H. pylori urease operon, ureABIEFGH, into the pRSETA vector to create the pHpA2H vector. We will introduce the mutations into the ureF gene in the pHpA2H vector. E. coli will be transformed with wild-type and mutant pHpA2H vectors, or the negative control plasmids (pHpAB and the empty vectors). The bacterial cells will be grown in the presence of 0.5 mM nickel sulfate, and were induced overnight with 0.4 mM IPTG. After cell lysis by sonication, urease activity of the bacterial lysate will be assayed in 50 mM HEPES buffer at pH 7.5 with 50 mM urea substrate, and will be measured by the amount of ammonia released using the method described in ref. [40].

If the dimerization of UreF and UreF/UreH is essential to the maturation of urease, the mutations that break the dimerization will also abolish the maturation of urease. On the other hand, if the maturation of urease is not affected by these mutations, it is likely that the dimerization is not essential to the urease maturation.

Our preliminary data showed that removal of the C-terminal residues of UreF breaks the UreF/UreH complex, and abolishes the maturation of urease. The availability of a preliminary structure of UreF/UreH complex allows us to introduce site-directed mutations that are designed to break the UreF/UreH interaction, and to further investigate the functional importance of UreF/UreH interaction. Our structure showed that upon complex formation, the C-terminal residues of UreF become structured and form an extra helix (helix-11) that dock to a binding cavity of UreH. Three hydrophobic residues V235, I239, and M242 on helix-11 are buried to a hydrophobic pocket of UreH.

To further investigate the functional importance of UreF/UreH interaction in urease maturation, we will create alanine and hydrophobic-to-polar (e.g. V?N) substitutions at V235, I239 and M242, which are designed to break the UreF/UreH complex formation. We will test if these mutations affect the formation of the UreF/UreH complex by the GST pull-down assay described in 3.1.1, and if they affect maturation of urease as described in 3.1.3. If the interaction between UreF and UreH is essential to the maturation of urease, we anticipate the mutations that break the UreF/UreH interaction will also abolish the maturation of urease.

The UreF/UreH is also known to form bigger complexes with UreG, and with the urease (UreA/UreB). How UreF/UreH complex associates with UreG, and the urease to form the pre-activation complex (UreA/UreB-UreG/UreF/UreH) for the maturation of urease is poorly understood. It has been reported that UreG does not interact directly to the urease, suggesting the UreF/UreH complex serves as a bridge that recruits UreG to the activation complex.

Our group has recently collected 2.5Å diffraction data for the H. pylori UreF/UreH, and has obtained a preliminary structure of the complex, which allows us to identify surface residues of UreF and UreH. To map the interacting surface of UreF/UreH complex for binding of UreG and the urease (UreA/UreB), we propose to perform alanine-scanning mutagenesis of surface residues on UreF and UreH. We will first focus on relatively more conserved surface residues (For UreF: P44, I45, Y48, S51, E55, Y72, E119, R121, Y183, K195, Q201, Q205, H244, E245, R250, L251, S254. For UreH: D60, G61, T78, K84, P111, I115, F177, E140, R146, E151, R213). We will also introduce multiple substitutions at these residue positions, if they are close in space according to the preliminary structure of UreF/UreH complex. We will first test if these mutations affect UreF/UreH interaction as described in 3.1.1. If so, we will exclude those mutants from the library.

After we have created the mutant library of UreF/UreH complex, we will test the mutants' ability to form complex with UreG, and with the urease (UreA/UreB). In brief, mutants of the GST-UreF/UreH complex will be co-expressed in E. coli. Our preliminary data suggest that the GST fusion tag will not interfere with binding of UreG or UreA/UreB (Fig. 4B). The bacterial lysate of GST-UreF/UreH (or its mutants) will be mixed with bacterial lysate expressing UreG or UreA/UreB, and then loaded to a GSTrap column for the pull-down assay. For those mutations that break the interaction, we will also perform the reciprocal pull-down in which the GST-tag is fused to the UreH, UreG, UreA or UreB. This is to confirm that the breakage of interaction is due to the mutations, but not due to a nearby GST-tag.

To address the question if the interaction between UreF/UreH and UreF (or UreA/UreB) is essential to maturation of urease, we will test the ability of the UreF/UreH mutants to activation urease in vivo as described in 3.1.3. If the interaction is essential to urease maturation, we anticipate the mutations that break the interaction will also abolish the urease maturation.

3.4.1 Expression and purification of UreG/UreF/UreH complex - We have established an efficient expression purification protocols for the ternary UreG/UreF/UreH complex. His-GST-tagged UreF, UreG and UreH will be co-expressed together in E. coli BL21(DE3) strain using the expression plasmids pET-Duet-HisGST-UreF/UreG and pRSF-UreH. After affinity chromatography purification, the His-GST fusion tag will be removed by the PreScission Protease (GE Healthcare). The protein complex will be further purified by gel filtration. Typical yield of the UreG/UreF/UreH complex is ~ 5mg per liter of bacterial culture.

3.4.2 Optimization of crystallization conditions - Preliminary screening of crystallization conditions was performed. We have already obtained crystals of the UreG/UreF/UreH that diffract to ~3Å (Fig. 8). We will further optimize the crystallization condition by grid-searching the pH and precipitant concentrations, and addition of additives or detergents. Quality of the diffraction data will be used to guide optimization of the crystallization conditions. We will also optimize the cryo-protection procedures (e.g. the choice of cryo-protectants and their concentration) to improve the quality of diffraction data collected. When necessary, we have access to synchrontron beam line at Diamond Light Source, Oxford, through collaboration with Dr. Yu-Wai Chen (King's College London).

3.4.3 Phase determination - We will first attempt to phase the structure by molecular replacement. At the time of writing this proposal, we are refining the structure of H. pylori UreF/UreH complex. We will use the UreF/UreH complex structure as a search template to solve the phase of the UreG/UreF/UreH complex by molecular replacement. In parallel, we will also prepare selenium-methionine labeled sample of UreG/UreF/UreH for multi-wavelength anomalous diffraction (MAD) phasing by expressing the protein complex in minimal medium containing Se-Met as described in Doublie [41]. The H. pylori UreG, UreF, and UreH proteins contain 9, 10, and 8 methionine out of 199, 254, and 265 residues, which should provide enough phasing power for MAD phasing. The PI's group has previously established the expression protocols for Se-Met labeling for H. pylori UreF, and determined its structure by MAD phasing. We have access to synchrontron beam line at Diamond Light Source for collection of MAD data.

3.4.4 Model building and refinement - Models will be built interactively by the program COOT [42], and refined using PHENIX [43]. The progress of refinement will be monitored by Rfree- and R-factors. Quality of the crystal structure will be validated by the program MOLPROBITY [44].

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The Mapping Of Erecta Genes In Arabidopsis Thaliana Biology Essay

Biology » The Mapping Of Erecta Genes In Arabidopsis Thaliana Biology Essay

Arabidopsis thaliana is used as a model organism for analyzing genetic and biochemical process in higher plants. Due to its small genome size (Approximately 100 Mb) and availability, Arabidopsis is used as a model plant for most of the experiments. In this experiment we have analyzed the function and position of ERECTA gene with respect to expression of trait and genotyping. Genotyping was carried out using PCR analysis. The mapping was done for the whole genome of Arabidopsis RIL’s using the indel markers. By using the recombinant inbred lines of Ler and Col the phenotype and genotype data were compared manually. Furthermore, the QTL analysis was used to identify the location of ERECTA gene on the chromosome along with the help of markers. From the results it is seen that the ERECTA gene is located at the second chromosome of Arabidopsis thaliana between the markers m220 and m251 (~between 22.6 cM and 37.8 cM). However marker m226 was also found to have minimum control on the expression of the trait (silique and pedicle length).

Key words: Arabidopsis thaliana, ERECTA, QTL, PCR, marker

Arabidopsis thaliana is a widely distributed plant which is also used as a model organism in plant science research. Arabidopsis thaliana has more than 700 natural accessions around the world. Among the ecotypes of Arabidopsis Ler and Col are the most common ecotypes which are used for genetic and molecular studies (Anderson and Mulligan, 1992). Among those Ler is isolated from mutagenized seed populations of Arabidopsis (Rédei, 1992) and it lacks ERECTA gene. The ERECTA gene controls many physiological processes during Arabidopsis plant development such as inflorescence, internodes and pedicel elongation and leaf and siliques morphogenesis (Scott J. Douglas, 2002). From the genetic control studies, the stem development with Arabidopsis mutation (erecta) has reduced internode length (Hanzawa et al., 1997) which leads to reduced plant height. These basic morphological characters are used to differentiate the ecotypes. In this experiment our main aim is to identify the position of the ERECTA gene and its linked marker in the genome (five chromosomes) of Arabidopsis. By using the ERECTA mutated ecotype Landsberg erecta and normal ecotype Columbia the Recombinant inbred lines are produced. The phenotyping (morphological character) and genotypic correlation of inbred lines is compared with the molecular markers and the location of the ERECTA is identified. The main application for this type of experiments is to identify the location of genetic factors (quantitative trait loci or QTLs) on the genome using molecular markers and used while making selection and breeding decisions to increase the selection efficiency of the trait.

The two Arabidopsis ecotypes Landsberg erecta (Ler) and Columbia (Col-0) were used as parents to form Recombinant inbred lines.

The initial cross with Ler and Col was done to form F1 generation. After that the F1 went 8 generation of recurrent inbreeding to increase homozygosity. The fig.1 shows the systematic diagram of RIL.

Fig 1. The recombinant inbred line population.

The different RIL samples were collected for DNA extraction. Collected plant samples were grinded in eppendorf tube by placing them in liquid nitrogen. 500 µl of extraction buffer was added and incubated at 650 c for 20 minutes. The extraction buffer contains 100 mM Tris (=Trizma base); M=121.14 g?mol-1, 1.4 M NaCl; M=58.442 g?mol-1, 20 mM EDTA (C10H16N2O8, M=292.24 or C10H14O8N2Na2·2H2O 372.24 g?mol-1), 2% v/v CTAB (N-Cetyl-N,N,N- trimethylammoniumbromide, M=364.46 g?mol-1). Then 500 µl of phenol- chloroform-isoamyl alcohol was added in the ratio of 25:24:1 and centrifuged at 13000 rpm for 5 minutes. The supernatant was transferred to another tube. Equal volume of chloroform- isoamyl alcohol was added in the ratio of 24:1 and centrifuged at 13000 rpm for 5 minutes. 300 µl of supernatant was transferred to a new tube. 30 µl of sodium acetate and 750 µl of 100% ethanol were added. The tubes were placed at -200 c for 10 minutes and centrifuged at 13000 rpm for 15 minutes. The ethanol was decanted. To the sample 175 µl of 70% ethanol was added and centrifuged at 13000 rpm for 5 minutes. The excess ethanol was removed using speed vacuum. The pellet was re-suspended in 50 µl of water. The concentration and purity of the DNA sample was measured on a Nanodrop.

The genotyping of Ler, Col and their recombinant inbred lines (00, 03, 11, 15, 21, 27 and 51) were analyzed by using the known insertion or deletion markers. The markers and the primers with their base pair expression are given in table 1.

m213

GCACCTCATGAAACCGATGCAAGT

ATCTTTGTTTGTGGTGGCAGAGCC

222

176

m251

GCGCACCTCTGTACAGTCTCT

CCTCTGGGTCAAACGAAGAA

477

445

ERECTA

ATCCCCAGCACGAATGTTTA

GGCAAACCAAAGAAAACCAA

1035

413

m220

TTGCGTCATGTGGTGACTCT

CGAGATTGAATGGTGATCCA

513

468

m457

GACCGGTCTTACATGACCAA

AACGGGTGACTTCTGGTTTG

616

533

m600

CTCGCAGTGGTGATGAAGAA

GCAGCTTGGTTCTGTGATGA

502

387

m555

AAAAGCAGAGAAGCAAAACACA

AGTTGGTGAAAGAGCGGCTA

537

310

Table1. The Markers and primers used for genotyping.

The 12x concentration of buffer, water, dNTP, polymerase and primers (Table 2.1) were added to a PCR tubes along with the DNA of different genotypes. The tubes were mixed well (without air bubbles) and placed in ice. Water is used as a negative control. The PCR program was set as shown in table 2.2. The tubes were placed in PCR machine and allowed to run for 35 cycles.

10x ThermoPol buffer

2.5 µl

30 µl

dNTP (10mM)

0.5 µl

6 µl

Taq polymerase (5U/µl)

0.2 µl

2.4 µl

Primer 1 (10µM)

0.6 µl

7.2 µl

Primer 2 (10µM)

0.6 µl

7.2 µl

DNA template

0.5 µl

H2O

20.1 µl

241 µl

96 °C

5 min

94 °C

30 sec

60 °C

30 sec

35 cycles

72 °C

1.5 min

72 °C

10 min

4 °C

hold

Table 2.1 The reagents for PCR Table 2.2, PCR program

5 µl of loading dye was added with the products obtained from PCR. The samples were then loaded into agarose gel. DNA ladders were added to the first and last lanes of the gel. The gel was run at 100 V for 1-2 hrs.

From the grown RIL and ecotypes of Arabidopsis the phenotyping was done. The morphological characters such as short plant, silique width, compact inflorescence and short silique length, pedicle length and petiole length contained plants were considered as landsberg erecta. For the Columbia ecotype tall plant, long silique length, pedicle length, petiole length, thin siliques width and with disperse inflorescence considered as phenotypic characters. The phenotypic data was compared with genotypic data obtained from the electrophoresis. Then the phenotypic data was compared with standard marker data and scoring was done.

The widely-used methods for detecting QTLs such as single-marker analysis, simple interval mapping (Liu, 1998) were used to find, whether the marker is linked to a QTL and the position of the QTL on the map.

The DNA of ecotypes and RIL were analyzed by using the insertion and deletion markers with the help of electrophoresis (Fig .2) and the data obtained was given in table 3. Based on the size of the DNA the ecotypes were differentiated as shown in fig 2. For DNA size refer table 1. For many of the markers there was no DNA band observed for the Parents and RIL’s.

Fig 2. The gel picture of Arabidopsis ecotypes and their RIL’s according to the marker.

From the table (Table 3) it’s clear that some of the markers such as m600, m555 and 457 were more different from the phenotype. Some of the other markers like m220, m213 and ERECTA don’t have clear genotypic results to compare with phenotype. However the marker m457 results have closely related to the phenotypic result. So there may be contamination or practical errors be occurred during the DNA analysis.

m213

Ler

Col

Ler

m251

Col

Ler

Ler

Col

Col

Col

Ler

Ler

Col

ERECTA

Ler

Ler

Ler

Ler

m220

Col

Col

m457

Col

Ler

Col

Col

Col

m600

Col

Ler

Col

Ler

Ler

Ler

m555

Col

Ler

Ler

Ler

Ler

Col

Col

Ler

Col

Phenotype

Col

Ler

Ler

Col

Col

Ler

Ler

Ler

Ler

Table3. The result form agarose gel electrophoresis and phenotyping.

The morphological characters such as type of inflorescence (compact or dispersed), plant height, silique length, silique width, pedicle length and petiole length was analyzed from the recombinant inbred lines of Ler and Col (Table 4). As it was said earlier (methods), based on the morphological characters, the ecotypes were differentiated and noted down. Form the result it was observed that most of the RIL’s expressed Ler phenotype.

CS/N1900

22

1

14

1

6

0

ler

CS/N1901

20

0

16

0

12

1

col

CS/N1903

23

0

18

0

13

0

col

CS/N1905

27.5

0

20

0

11

1

col

CS/N1910

18.5

1

12

1

3

0

ler

CS/N1911

32.5

0

14

0

10

1

col

CS/N1913

8.5

1

6

1

6

0

ler

CS/N1915

18.5

1

15

0

6

1

ler

CS/N1919

35

0

18

0

12

1

col

CS/N1921

13.5

1

9

1

4

0

ler

CS/N1923

20

1

10

1

6

0

ler

CS/N1924

32.5

0

17

0

11

1

col

CS/N1927

16

1

10

1

9

0

ler

CS/N1929

28

0

16

0

13

1

col

CS/N1933

9

1

6

1

3

0

ler

CS/N1934

20

1

10

1

6

1

ler

CS/N1935

20

1

11

1

5

0

ler

CS/N1937

23

1

9

1

5

0

ler

CS/N1938

24.5

1

15

0

4

0

ler

CS/N1942

23.5

1

10

1

4

0

ler

CS/N1945

34

0

16

0

11

1

col

CS/N1946

22.5

0

15

0

10

1

col

CS/N1948

30

0

15

0

11

1

col

CS/N1951

5

1

5

1

2

0

ler

CS/N1953

25

0

16

0

10

1

col

CS/N1954

35

0

18

0

13

1

col

CS/N1957

15

1

11

1

5

0

ler

CS/N1958

17

1

10

1

8

0

ler

CS/N1959

18.5

1

14

1

5

0

ler

CS/N1960

18.5

1

12

1

6

0

ler

CS/N1963

21

1

11

1

8

0

ler

CS/N1966

7

1

10

1

8

0

ler

CS/N1969

12

1

10

1

3

0

ler

CS/N1971

38.5

0

16

0

8

1

col

CS/N1974

25.5

1

11

1

3

0

ler

CS/N1975

11.5

1

8

1

6

0

ler

CS/N1978

36.5

0

19

0

13

1

col

CS/N1980

16.5

1

9

1

6

0

ler

CS/N1984

12.5

1

7

1

8

0

ler

CS/N1985

12

1

12

1

4

0

ler

CS/N1988

13

1

4

1

6

0

ler

CS/N1989

34

0

17

0

14

1

col

CS/N1990

23.5

1

14

1

3

1

ler

Table4. The morphological characters of RIL of Ler and Col. In inflorescence 1 denote compact inflorescence and 0 denote disperse inflorescence. For leaf shape 0 and 1 indicates presence and obscene respectively. Like that, Silique length was also denoted by 0 and 1.

The genotype of the markers was scored with the help of phenotypic data (Table 5). In the table the phenotype observed was compared with the genotype data, by highlighting the similarity between phenotype and genotype. Form the table5 marker m220 had highest value of 37 followed by m251with 36 and m216 & m326 respectively with 29.

Table5. The data of scored marker with observed phenotypic data of RIL’s. Highlighting indicates genotype and phenotype were matching.

The QTL results based on single marker analysis and interval mapping are shown in Fig.3.1, 3.2, 3.3, 3.4. The QTL analysis for height, silique and pedicle length indicated that (Fig.3.1, 3.2 & 3.3) marker m220 had peak above the threshold level. For silique length and pedicle length the QTL showed (Fig 3.2 & 3.3) two peaks (high and low) above the threshold level for the markers m220 and m226 respectively.

Fig. 3) 3.1.The QTL of plant height with peak on m220 (indicated by arrow, 3.2 & 3.3, QTL of silique length and petiole length with two peaks above threshold level on m220 and m226 and 3.4, the marker map of Arabidopsis showing the place of ERECTA gene.

The genotype result gave an idea to eliminate the marker which was not closely related to phenotype such as m600, m555 and 457. But, the genotype result did not given clear conclusion about the marker that closely related to ERECTA gene. The genotype data of m220, m213, m457 and ERECTA did not express DNA band for most of the RILs (Fig. 2). However, the score obtained from the comparison of phenotype with standard genotypic data provide some evidence of the location of ERECTA gene when compared with molecular map of Arabidopsis thaliana. It showed that m220, m251, m216 and m226 were closely related to the ERECTA gene. Still we cannot say clearly that ERECTA is located somewhere between m220, m251, m216 and m226 because the marker m226 and m326 were located at third chromosome of Arabidopsis thaliana. Additionally, the markers m326 and m226 showed some effects on silique and pedicle length which were observed in both QTL analyses and RILs scoring. So it can be said that these genes also have a little effect on controlling the expression of silique and pedicle length of Arabidopsis thaliana. By using QTL analysis the finest details about location of ERECTA gene was obtained. From the QTL data it was clearly shown that the ERECTA gene is located between m251 and m220 (Fig. 3.4).

We thank Tom martin, Jonas Ross and Luisa Ghelardini for providing technical support and assessment during the experiment.

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The Distance Slaters Travel In Different Temperatures Biology Essay

Biology » The Distance Slaters Travel In Different Temperatures Biology Essay

Introduction: The Slater Porcellio Scaber is an Arthropod and is a member of the biological class Crustacea. The Slater has a flattened segmented body and 7 pairs of legs. The Slater is nocturnal and feeds on dead plant matter and vegetables. The Slaters I collected were found in dark and damp places with a temperature of 17.4 degrees Celsius in a gully behind bark on rotting trees and under rocks. The Slaters I found were clumped together as this is a method of conserving water in warmer temperatures or in regions of low humidity. Water loss can lead to desiccation as can the exposure to light so the Slaters I found were unexposed to light. This is due to Slaters being negative photo taxis as exposed to light Slaters can suffer desiccation (“Desiccation is the state of extreme dryness, or the process of extreme drying” quote from http://en.wikipedia.org/wiki/Desiccation.) This is because Slaters do not have a waterproof waxy cuticle to keep water/moisture inside there body. Also Slaters lose more water than other Arthropods because their lungs are located on the abdomen and during respiration water is a product which can be lost via a small pore on the outside of the body. Due to losing water more rapidly than other Arthropods Slaters have physical and behavioural adaptations to maximise and maintain their water intake in order to avoid desiccation.

A structural adaptation Slaters have is a pair of uropods. Slaters are able to absorb water through uropods. The uropods are located on the posterior of the insect. To obtain the water the Slater will squeeze the uropods together and touch it against the surface with water on it. Capillaries located inside the uropods ‘suck’ the water up and into the anus.

Slaters are able to absorb water directly through their exoskeleton. However this is only possible in regions of high humidity.

A behavioural adaptation is in hot or dry conditions the Slater is able to release a body odour that attracts other Slaters. The Slaters then clump together as a form of defence to prevent loss of water. They do this because they need to reduce their surface area and for a lone Slater this is not possible. Reducing surface area means reducing the area in which water is lost via diffusion through the exoskeleton due to the lack of the waxy cuticle.

Slaters are nocturnal. Although this is also to avoid predators such as birds, frogs, some beetles and some spiders it is also to avoid sunlight which can lead to water loss and desiccation. Also at night the temperature would be a lot cooler than during the day. Slaters may have developed this behavioural adaptation because being active during the day would lead to significantly more water loss than when they are active at night. A large loss of water can lead to desiccation.

My investigation is based on whether the temperature in which the Slaters live in naturally benefits their survival or not. As warmer temperatures can lead to water loss and for Slaters this can lead to desiccation. I have designed an experiment to measure the activity levels of Slaters by measuring the distance they travel (in cm) at different temperatures (5 degrees Celsius, 10 degrees Celsius, 15 degrees Celsius, 20 degrees Celsius, 25 degrees Celsius and 30 degrees Celsius). The results from this experiment will allow me to determine whether or not Slaters have increased or decreased activity levels in warmer or cooler temperatures. Results will allow me to make a conclusion on whether the temperature in which they live in naturally plays a vital role in the survival of Slater populations. I would expect to see in my experiment Slaters covering a larger amount of distance in the warmer temperatures. I am expecting to observe this behaviour because the Slaters will be trying to avoid these warmer temperatures in order to prevent water loss which could then lead to desiccation.

Aim: To determine whether Slaters (Porcellio Scaber) have an increased/or decreased activity levels in different temperatures measure by the distance they travel in a set amount of time. Using the results make a conclusion on whether temperature plays a vital role in the Slaters survival as a population in their natural environment by reducing the risk of desiccation.

Hypothesis: I think the Slaters will cover more distance in the warmer temperatures (e.g. 20°C, 25°C and 30°C) as an attempt to find an escape to a cooler temperature. This is because in warmer temperatures more water is lost and excessive water loss can lead to desiccation. I think that in the cooler temperatures the Slaters will not cover as much distance because they will not be trying to avoid the temperature as desiccation would occur at a significantly slower rate in colder temperatures.

Equipment List:

Lux meter

Data logger with humidity measure attachment

Ice cream container

Thermometer

Stop watch

Sharpie

String

Boiling water (depends on the temperature you are trialling)

Ice water (depends on the temperature you are trialling)

200 similar sized Slaters (Porcellio Scaber)

Paper (to line the bottom of the ice cream container with)

Cello tape

Question: To measure how temperature affects Slaters activity levels by the distance they travel in a set time. With the results make a conclusion on whether the temperature in which they live in naturally benefits their survival.

Gather 200 similar sized Slaters. Slaters need to be of similar size as larger Slaters may be able to cover further distance than smaller Slaters are able to. Keep Slaters in a tank outside (out of direct sunlight) that replicates their natural environment. Also during the experiment we want to replicate their natural environment as much as possible (e.g. light intensity and humidity). This is to ensure the Slaters respond to the changes in temperature and not other factors.

Collect a sharpie, string, measuring instrument, thermometer, lux meter, humidity meter, boiling water( depends on the temperature), ice water (depends on the temperature), data logger, ice cream container.

Line the bottom of an ice cream container with a sheet of white paper. Ensure you use the same size container for all of the trials during your testing. As a change in size may change the Slaters movement and then it would not be a fair test. Also ensure container is same height as if the container were to be not as high for some temperatures in your experiment the Slater may spend the 2minutes trying to climb out and not moving around the container in response to the temperature.

When carrying out the experiment make the room as dimly lit as possible. This is to prevent the Slaters from moving in response to bright light and not temperature. As Slaters are negative photo taxis (move away from light) having the experiment in a bright room will cause the Slaters to move in response to the light as they will be trying to move away from it. Light is a controlled variable so measure the light intensity using a lux meter before every trial and every change in temperature. Take 5 readings and calculate an average. You must take 5 readings to obtain the average as light intensity is constantly changing so by taking 5 readings and calculating an average you will be getting a more accurate light intensity measure rather than if you were to just use one reading as your light intensity. Also the humidity of the room is another controlled variable. Measure humidity using a data logger with a humidity measurer attachment after every trial and change in temp just to ensure it remains fairly constant (within 10% difference in the humidity readings). We must keep the humidity level as constant as possible as different humidity levels may cause the Slaters to act differently and we want a response form the Slaters due to temperature to which they are exposed to and not humidity levels.

To obtain cooler temperatures (e.g. 5 and 10 degrees Celsius) cool down the inside of the ice cream container by placing it in a water bath containing ice water. Once the temperature has reached the desired coolness and has remained at the desired temperature for 5minutes you are ready to start testing. Measure temperature using a thermometer. Ensuring the temperature maintains the same for 5minutes means when you start the testing it won’t start changing. This is important because if the temperature were to increase or decrease during the testing the Slater would respond to the change in temperature and not to the original temperature which is the one we are testing the Slaters activity levels on.

Place a randomly picked Slater in the ice cream container. (Do not get the Slater from the tank you are keeping it in until you are ready to start the testing). Measure the Slater using a ruler to check it is between 1-1.5cm. The Slaters used for testing must all be between 1-1.5cm because if you were to use smaller Slaters and larger Slaters for the experiment you may receive different results as the larger Slaters may be able to move faster so they could cover a larger distance in the 2minutes. Wait for 2minutes to allow the Slater to adjust to its new environment. If you were to start timing straight away the Slaters response may not be due to the temperature but due to its new surroundings.

After the 2minutes ensure the temperature is still constant. Now start a timer for 2minutes and as the Slater moves around the ice cream container draw a line behind it and line the path it takes. Make sure you do not follow the Slater with the sharpie too closely as it may respond to the sharpie following it and not the temperature. Leave about a 10cm gap between sharpie and the Slater.

Once the 2minutes is up remove the Slater and place in a separate tank to the other Slaters that replicates its natural environment. We don’t want to use the same Slater for the trials as this would not be a large enough sample size to represent a population.

Using a piece of string follow the lines (the path the Slater took) around the piece of paper that lined the bottom of the ice cream container. You may need to use cello tape to keep the string in place along the Slaters path. Once you have the length of string with the distance the Slater travelled in the 2minutes measure it using a ruler and record in a results table. A large amount of distance=high activity levels. A low amount of distance=low activity levels.

Now you are going to repeat the trial again so measure the temperature to ensure it is still constant. You may have to add more ice to the water bath to cool it down to the desired temperature again. Once the temperature is constant again and has remained constant for at least 5minutes you are able to repeat the trial again for 9 more Slaters. Each temperature you test you must have a sample size of Slaters of at least 10. This is because a large sample size will mean more accurate and reliable results that can represent a whole population of Slaters.

For each different temperature you test (5, 10, 15, 20, 25 and 30 degrees Celsius) repeat the same method but to heat up the inside of the ice cream container place the container in a water bath full of hot/boiling water and have the room well heated.

For each temperature use 10 Slaters and each temperature must be trialled 3 times (so 30 similar size Slaters will be used per temperature overall). This is to ensure accurate and reliable results as the sample size must be able to represent a whole population.

Record all results in a table. Remember between each trial and sets of testing record light intensity and humidity to make sure the two controlled variables remain as constant at possible. Also remember to measure temperature before each Slater is tested to make sure it is still at the desired temperature.

With results work out averages from each temperature (using the averages from the 3 trials per temperature). Use the averages to make a conclusion whether Slaters benefit their survival in their natural environment by living in the temperature they live in.

Light intensity: You must maintain a constant low light intensity throughout the testing. Because Slaters are negative photo taxis we want the room as dark as possible whilst performing the experiment so the Slaters respond to the temperature and not to the light intensity.

Humidity: You must maintain a constant humidity reading throughout the testing. You want the Slaters to respond to temperature and not humidity levels.

Perform testing at roughly the same time every day: Because Slaters are nocturnal we must perform the testing if done on different days at roughly the same time. If you were to perform some trials during the evening this is when the Slaters are normally active so if you had done the rest of the tests during the day your results would not be accurate.

Perform testing in same container: Make sure the container is the same length; width and height (preferably keep it the same container throughout the whole testing process). If the container were to change the Slaters may act differently. If the container were to be not as high as the original the Slaters may spend the 2minutes trying to climb out the container and not moving around the container in response to temperature.

Use similar sized Slaters for the testing and the same type of Slater: Make sure all the Slaters you use in your testing are between 1-1.5cm in length. Check this by measuring them with a ruler before you start the testing. The Slaters used for testing must all be between 1-1.5cm because if you were to use smaller Slaters and larger Slaters for the experiment you may receive different results as the larger Slaters may be able to move faster so they could cover a larger distance in the 2minutes. Also make sure you are using the right type of Slater as there are many different types found in New Zealand. You are using Porcellio Scaber, this Slater cannot roll into a ball and it is a little bit blue in colour so it is easily recognisable.

Temperature

Average Distance in cm the Slaters Travelled in the 2 minutes.

5°C

10°C

15°C

20°C

25°C

30°C

1.92cm

11.98cm

23.08cm

65.20cm

83.21cm

95.86cm

As seen on the table the Slaters travelled a very little amount of distance in the 5°C with an average distance of 1.92cm and as the temperature increased so did the amount of distance the Slaters travelled. All the measurements of distance shown are final averages from the 3 trials per temperature.

Conclusion: My aim was to determine whether Slaters have an increased/or decreased activity levels in different temperatures by the distance they travel in a set amount of time. In conclusion from the results of my experiment it shows that as the temperature increases so does the Slaters movement. This is shown by the distance in cm Slaters travelled in the 2minutes. For 5°C the Slaters travelled an average distance of 1.92cm. In 10°C the Slaters travelled an average distance of 11.98cm. In 15°C the Slaters travelled an average distance of 23.08cm. In 20°C the Slaters travelled an average distance of 65.20cm. In 25°C the Slaters travelled an average distance of 83.21cm and in 30°C the Slaters travelled an average distance of 95.86cm. Therefore my hypothesis was correct with my assumption that the Slaters would cover more distance in the warmer temperatures.

Discussion: The purpose of my investigation was to determine whether Porcellio Scaber have increased/or decreased activity levels in different temperatures. A trend was seen in the results I obtained. As the temperature increased by 5°C the Slaters movement increased by 10-20cm with the exception of the temperature change from 15°C to 20°C. The difference of the Slaters movement between the two temperatures was 42.12cm. From the results of my experiment it was clear the Slaters preferred the temperatures between 10°C and 15°C. 5°C was too cold for the Slaters and they moved a very minimal distance (average of 1.92cm). This is because as Slaters are cold blooded and are not self heating so they take in the temperature of their environment. In the cold temperatures Slaters cannot move quickly as particles are denser and thus move slower and enzymes cannot catalyze reactions as quickly in the cooler temperatures. If the Slaters were to live in a cold environment such as 5°C they would struggle to move especially away from predators and it would take them longer to find food as they could not travel as fast. This would be a disadvantage to the survival of Slaters as they would be more likely caught and eaten by predators and they may not be able to find enough food for them to survive as they would be moving very slowly. The other extreme is in the warmer temperatures the Slaters covered a large amount of distance. In 5°C on average the Slaters travelled 1.92cm and in 30°C on average the Slaters travelled 95.86cm. The difference of 93.94cm between the two temperatures for the Slaters movement is a significant difference. In the warmer temperatures (e.g. 20°C, 25°C and 30°C) the Slaters all covered a large distance. This is due to the Slaters trying to find an area that is cooler to escape the heat. Heat causes particles to move more rapidly as they become less dense. Therefore diffusion occurs at a more rapid rate in warmer temperatures and Slaters lose water via diffusion through their exoskeleton. This can then lead to desiccation. Desiccation is the dryness resulting from the removal of water. Slaters are more prone to desiccation than other arthropods as they lack a waxy cuticle on their exoskeleton. The waxy cuticle which is found on other arthropods prevents water loss. Slaters also lose water through respiration as water is a product of the chemical reaction. Also the Slaters lungs which are located on the abdomen open to the outside of the body via a pore. This means Slaters will lose a greater amount of water from their respiratory surfaces than other Arthropods. From my experiment part of my aim was to find out whether temperature plays a vital role in the Slaters survival as a population in their natural environment. From my results of my experiment I have come to the conclusion that temperature does play a vital role in the survival of Slater populations. Optimum temperature for Slaters would be around 15°C because the Slaters were still able to move at a rate and cover a suitable amount of distance (23.08cm) in the 2minutes that would allow them to escape from predators. At around 15°C the Slaters would not have to be concerned of excessive water loss which could lead to desiccation. The jump of the distance travelled by the Slaters between 15°C and 20°C (42.12cm) shows that when the temperature is around 20°C the Slaters start trying to escape and search for an area cooler in temperature so this would not be an ideal temperature as they would be prone to desiccation in this temperature.

Evaluation: My results and conclusion from my experiment are valid because I made sure my method was reliable by controlling any factors that the Slaters could respond to if they were to change. The steps I took to make my experiment a fair test to ensure valid results are:

When I collected the Slaters from the gully and put them into a tank I made sure that the tank replicated their natural environment as much as possible by placing pieces of bark off the tree I found them on in the tank. I also added leaves, dirt and other plant debris. Also each day I added some water into one corner of the tank and also gave the Slaters vegetables such as potatoes to eat. The reason I kept the tank the Slaters were in as similar to their natural environment as possible and fed them is because I did not want the Slaters to become stressed or deprived of food and water as this could have significantly changed my results.

During the testing I controlled light intensity by measuring the light intensity using a lux meter after every trial and before every change in temperature. I also kept the room as dark as possible because Slaters are negative photo taxis which means they move away from light. If I were to perform the testing in a room well lit up the Slaters would respond to the light and temperature not just the temperature which is the only factor we want the Slaters to be responding to in order to receive valid results.

I also controlled humidity levels by measuring humidity before after every trial and before every change in temperature to ensure it remains relatively constant. I tested the humidity levels using a data logger with a humidity reader attachment. The reason for controlling humidity is if the humidity were to change for different temperatures the Slaters may respond to the humidity level but temperature is the only factor we want the Slater to be reacting to.

I performed the test at roughly the same time of the day if I was doing trials on different days. I did this because Slaters are nocturnal which means they are active at night. If I were to perform the testing in the evening the Slaters would already be more active in their natural environment so than if I were to perform the testing during the day so this could cause my results to be unreliable. To eliminate this from happening I ensured that I performed the testing midday-early afternoon as this means that all the Slaters I would have used for testing would be displaying the same activity levels at that time of the day in their natural environment.

During my experiment I used the same size container to perform the testing in. This is important because if I were to change the height of the container the Slaters may spend more time trying to climb out than moving around the container in response to temperature. In the bottom of my container I made sure I used the same surface for all the trials (sheet of plain white paper). If I were to not use paper on the bottom the Slaters may find the surface slippery and not be able to move naturally which could lead to inaccurate results.

During my testing I made sure that when I was dotting/lining the path the Slaters took around the bottom of the container I did not follow them too closely with the pen. I left around a 10cm gap between the sharpie and the Slater. I did this because if I were to line the path the Slater took directly behind the Slater as it moved the Slater may become frightened and try to avoid the pen so it would be moving in response to the pen behind it following too closely and not the temperature.

Another way of ensuring I obtained valid results from my experiment is if I started testing but found the Slater was acting abnormally I did not use the results from that experiment. If I were to use the results from the trial it may make my mean a lot smaller or a lot larger than what it really should be. An example of not using Slaters that act abnormally is I started to test one of the Slaters in 25°C and very quickly I noticed it had 2 legs missing which affected they way it moved and it was not covering the same amount of distance all the other Slaters had in 25°C. So I placed the Slater back into the set up tank and chose another Slater to use for the testing. If I were to use this Slaters results it may have brought down the average distance travelled for 25°C.

The Slaters I used for the testing were all between 1-1.5cm in length. I measured the Slaters I used for the testing because I had to make sure they were all similar in length. This is because if I were to use a large Slater for one test it may be able to cover a larger distance than smaller Slaters are able to. Having the same sized Slaters means that in a constant temperature they should all be able to cover equal amount of distance.

Another way of ensuring I had valid results was I made sure the Slaters got used to the environment before I started testing. I did this by leaving the Slater in the cooled down or heated up ice cream container for 2minutes before I started timing and recording its movement. I did this because if I were suddenly to grab a Slater from its natural environment and place it in an ice cream container and start timing straight away the Slater may respond to all the factors in its new surroundings and not to just the temperature.

For each temperature I tested 10 Slaters and I then repeated that 2 more times so in total I used 30 Slaters per temperature over 3 trials per temperature. So overall I used 180 Slaters for my experiment. I used such a large sample size because I needed to perform a test with enough Slaters so that it could represent a population of Slaters. If I had not of used such a large sample size I may have had inaccurate results as only a few Slaters cannot represent a whole population. If I had used a few Slaters they may have had a disease or not be acting normally due to a change in environment that consists of far less Slaters than what they were used to. This would of affected their natural behaviour because they would not be able to clump together to reduce water loss as there may not be enough Slaters for this to work. If I had then chosen to go ahead with my experiment using only a small sample size my results would be invalid and not be able to represent a whole Slater population. I also repeated my trial per temperature 3 times. I repeated my trials 3 times per temperature using 10 Slaters per trial because repeating the experiment multiple times verifies that your results are accurate and consistent. If I had only used 10 Slaters per temperature and not repeated this 2 more times I may have inaccurate results because some of the Slaters used may have displayed unusual reactions to the temperatures which would have resulted in my final averages being inaccurate. But because of having a large sample size and trialling the experiment 3 times if one Slater had not moved as far or too far in the 2minutes it would not affect the averages or have very minimal effect.

Because of all the precautions I took to ensure my test was a fair test I am positive my results are valid which lead to a valid conclusion.

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