In the florescence technology, the sample under study is itself the source of light. Florescence technology microscopes imaging depends on the phenomenon of light energy emission occurring in certain materials, in which material itself emits energy. This energy can be detected as visible light with specific wavelength. florescence in components of sample can be detected through treating sample with florescence chemicals, or even some cellular components are florescence themselves, such as chlorophyll.(1)
Applying florescence to the sample
When samples are studied, florescence material emits long wave lengths while non florescence substances (background) remains dark. In order to obtain a florescence image, florescence dyes have to be attached to the sample required to be studied. Florescence dyes are called fluorophores and they can be attached to the specimen by two ways. Firstly, is through florescence staining, through which a staining fluoresce dyes are applied to the sample. The other way, is through immunofluoresence techniques, in which fluorophores are located in antibodies that have to be attached to the sample by immunological reactions.(2)
What is fluorophore?
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It is the substance that labels the desired studied sample under microscope, in order to illuminate it so that it can be clearly detected. After fluorophore is attached to the sample, they absorb illumination light and emits a longer energy wave light. Specific filters with specific wavelengths can be used to filters light and view clearly only what is fluorescence.(3)
How does florescence microscopes work?
Florescence microscopes operates through allowing light, emerging from the microscope's light source, to excite florescence material in the sample, and thus, the image is formed. This can be explained in the following steps: First, florescence filters in the microscope passes florescence light resembling the sample's florescent material. Secondly, this light radiation hits the atoms of the florescence sample and electrons of sample are excited to a higher energy level. Afterwards, electrons loses their excited energy and return back to their original energy level emitting light photons. The light emitted is separated from the much bright excitation light through a second filter. It is important to mention that emitted light has a lower energy and a longer wavelength than original light used. Finally, florescence areas will be visualized and images are interpreted in attached computers containing specific softwares (Figure 1).(1,3)
Figure 1: Fluorescence: When electrons gains energy, they are excited from ground state to higher energy level state. Afterwards, they lose the energy returning back to ground state and emitting a photon. (own picture (no reference- copyrighted))
Uses of florescence
Florescence technique is progressively used in field of science, either the medical or the biological ones. It enables study of cells and various cellular components and mechanisms with increasing details and specificity.. For instance, studying certain antibodies, cluster of differentiations , cellular markers markers, certain antibodies, cellular signals. (1) Additionally, it is used to study apoptosis and perform it viability studies on cell to determine their living status and if they are living or dead. Florescence is applied in studies concerned with examination of genetic material in the cell (DNA and RNA). Other essential applications in floresence microscopy is also the using of conventional immuno-fluorescence technique in determining infectious diseases, or using fluorescence in situ hybridization (FISH) which is a novel and advanced application for florescence microscopy. TheÂ FISH is a florescent microscopy application in which there is direct localization of genesÂ and other DNA/RNA sequences in chromosomes or tissue, for instance, Turner syndrome karyotyping, Down syndrome karyotyping and any other karyotyping either prenatal or antenatal. FISH enables striking advances in experimental researches as well as clinical identification of diseases either prenatal or antenatal. Finally florescence microscopy can also be used in whileÂ and there is also studies of comparative genomic hybridization (CGH). The CGH is an application to detect genome and gene changes. It is a method of screening, that offer important information, especially in the field of for tumor pathology, and it gives information for the non-balanced genetic changes in the DNA under examination. (2, 3)
Problems in Florescence microscopes
Some problems revealed on the use of florescence techniques. Most importantly, the photo-bleaching and the photo-damage phenomena's. Photo-bleaching is defined as fading of florescence activity due to long exposure to the light illumination source. To lessen photo-bleaching, florescence microscopy devices are combined with other techniques that do not cause wasting of the fluorochromes, for instance, as in differential interference contrast (DIC) microscopy techniques and Hoffman modulation contrast (HMC) technique, Dark field illumination microscopes and phase contrast microscopes. (4) Other important problem is the elaboration of heat that can cause damage to the sample. Heat can be eliminated either from the complicated microcopy devices itself, or it can be causes due to insufficient cooling, such as cooling system inside the microscopy room. Bleed through , or so called cross over, is a problem that can be seen in wide field and laser scanning confocal fluorescence microcopies. In bleed through phenomenon, overlap in spectra of light can occur due to detection of a fluorophore through another channel and not it's specific channel of detection. (5)
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There are variable types of florescence proteins to be used in florescence research. Examples of florescent proteins are: GFP (green florescent protein) that comes from Aquorea, and it's available mutants, namely, YFP (yellow florescent protein) , RFP (red florescent protein) and CFP (cyan florescent protein).(7)
Uses of florescent proteins
Florescence proteins are used in various applications in florescence techniques. They can be applied for imaging cellular organelles, and can be used in molecular biology techniques such as florescence gene encoding. This have great impact in studying genes.(6)
Fluorescence is an pivotal technique in biomedical research. It allow visualization of various types of samples through light emitted from them when stained.
3- Bradbury, S. and Evennett, P.,Â Fluorescence microscopy, Contrast Techniques in Light Microscopy., BIOS Scientific Publishers, Ltd., Oxford, United Kingdom (1996).)
Live Cell Imaging
Definition of live cell imaging
Live cell imaging research is the studying of the living cells and living cellular components, behavior, structures and interactions through different imaging facilities, namely, advanced microscopes and high content screening systems. Live cell imaging is applied in science to achieve much clear views of cells as well as knowing more about their properties and biological functions through studying cellular dynamics. In the recent era, these technologies becomes widely growing and new advances and inventions are added every day. Accordingly , they became much more accessible, and accordingly, added a lot of capabilities to exploit areas of biology that could not be accessed before. It became one of the (of choice ) techniques in many areas of biology such as cell biology, neural biology, developmental biology, cancer biology, and various applications in biomedical research. (Figure 1) (1, 2)
UGENWebsitepictures2010GEN15_Sep0110LippLiveCellImagingLifeTech_Live_HeLaCells9239164772 Figure 1: Live cell imaging (2)
Techniques for live cell imaging
Many microscopic techniques are involoved nodays in living cell imaging. It became accessible to choose among wide range of such imaging techniques serving this area. For instance, and basically, confocal microscopy, that is the basic device for many live cell facilities, as well as it can view moving non fixated cells through pinhole optical plan imaging, depth cell imaging , and three dimensional as well as four dimensional cell imaging in which the fourth dimension is the time. Two photon microscopy is more advantageous than confocal in the deep intra-vital cell imaging, however, confocal is much beneficial in high detailed images. The use of two photon microscopy has made striking advances in intravital imaging of the living cells and detecting their interactions inside the body as well as forming clear three dimensional images to them (3,4) . Other confocal based devices includes; Spinning disc microscope which is optimal in acquiring life time view of what is happening inside the cells and cellular dynamics, FRET (Förster resonance energy transfer) which is used for studying of dynamic protein interactions, fluorescence recovery after photo-bleaching (FRAP) technique that is usually used in visualizing for monitoring protein or vesicle trafficking through attaching a florescence protein (usually GFP) is attached to the protein interested to be studied. There is also Total Internal Reflection (TIRF) microscopy, which is very convenient in experiment studying events located inside or close to cellular the plasma membrane of a cell. Stimulated emission depletion (STED) microscopy that, was discovered by Stefan Hell, enables creating images to structures and components beyond the limit of optical resolution. It enable creation of precise and clear images for such components. The STED operates through stimulation emission depletion mechanism through the use fluorescence dyes. Cellular growth, cell aggregation and cell movement can be efficiently observed using compound microscopes and contrasting methods, for instance, phase contrast microscopy and differential interference contrast (DIC). Moreover, larger specimens experiments such as viewing developing zebra-fish embryos, is commonly seen using stereo microscopes and microcopies. Fluorescence techniques became very advanced and gaining much more importance every day. Ion imaging is made now by using either special fluorescent dyes or proteins. On calcium binding to the ion channels , the special dyes and proteins change their emission behavior, and accordingly, facilitates studying them. (4)
Method of cell preparation
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Cells in experiments are of two types; they are either adherent or suspension cells.
For cell imaging cells are plated in petri dishes with coverslip. High quality coverslips should only be used, with cleaning them with strong acid or alkali and directly wash with water afterwards. The plastic coverslips have to be avoided in order to avoid auto-fluorescence and birefringence that alters imaging modes. Concerning culture chamber, it has to be large in respect to its lateral dimensions to be able to accommodate sufficiently the cells must be large enough in its lateral dimensions to contain sufficient cells to ensure comprehensive sampling of the population. About the depth of the medium surrounding the cells, it should be minimized as possible to guarantee high level of optical quality and avoid unclear images.(5, 6)
Factors affecting cells
Cells should be ensured to be alive and healthy, and they should be stored in a desirable manner. Strict control of the surrounding environmental factors is very essential in live cell imaging process. When cells are in microscopic stage area, many factors can lead to success or fail of the experiment. These factors includes: Firstly, physical parameters of the chamber containing the cells. Secondly, efficiency of cooling down systems and proper temperature control. The temperature emitted from the imaging device itself is very high that it can easily destroy the sample under examination. One of the other factors that should carefully be monitored are atmospheric conditions, namely, gas mixture and humidity. In addition, osmolarity and nutritional supplements should be cared for, in order to avoid losing cells. Finally and importantly, controlling photo-bleaching and photosensitivity resulting from the light source of the imaging device used is strikingly important. It has been advised to discard unhealthy cells and avoid imaging them for better results.(5, 7)
Possible artifact in live cell imaging is due to focus drift
Despite of recent advantages in optical microcopies, still there are some problems that put some obstacles in research procedures. An important problem in this topic is focus drift. Focus drift is defined as inability of the microscope to maintain the desired focal plane over sustained period of time. This artifact is not related to the natural movement of living cells required to be examined. Focus drift is mainly affected by temperature drifting problem called thermal drift. Thermal drift, which is considered to be the most common source of focal drift, is influenced by the huge amount of heat produced from the microscope itself. in addition to heat emerging from lab equipments and air conditioning. To solve this, high efficient cooling systems should be considered. Other important cause of focal drift is surrounding vibrations and movements. There are also coverslip flex artifacts, high magnification powers artifacts , and aperture oil emersion objectives.(8)
Live cell imaging is important in biomedical research in order to study different mechanisms that happens in living cells and tissues. Many devices nowadays has emerged for such purpose. Most importantly, confocal microscope, 2 photon microscope, spinning disc microscope and others.
3- Turku Bioimaging (More Than You Can Imagine), 2011
Introduction and pinhole idea
Confocal microscope is based on the idea of (pine hole) which allows light coming only coming from focal plan to reach the detector and discard other (out of focus) waves above and below. Its advantage over the conventional optical microscopes is ability to visualize deeper tissues, creating optical focal planes and interpreting them in three dimensional structure. This made the confocal to be increasingly and essentially used in cell biology researches. (1, 2,3)
Image formation in confocal microscopy:
Confocol microscope is a kind of optical microscope, through which image is formed through a pinhole. This pinhole is the pivotal component in the confocal image formation process. light source of confocal microscope is laser beam. This pinhole structure is located in the confocal plan of confocal microscope, while the specimen is located on the focal plan of confocal microscope. Focal and confocal planes are located on opposite sides with the lens between them First, image of confocal, located in the focal plan, is is illuminated by the lase beam which excites the florescence specimen. This laser scan is processed in a point by point manner through over the whole specimen plan. Afterwards, light is detected through a photon multiplier tube. Thereafter, detected light will pass through the pinhole. The importance of pinhole structure is to eliminate most of the light above and below the focal plane of the specimen. Accordingly, optical section can be created to the specimen. Afterwards, focus can be changed to create more and more optical sections to the examined specimen . Finally, all the acquired optical section from the sample can be collected together to form a three dimensional image of the sample, and this a great advantage that cannot be achieved by a conventional two dimensional microscope. Sectioning in confocal is optical, non invasive and comparatively fast, and accordingly, image also can acquired in a four dimensional manner, by which the fourth dimension is the time. Special softwares are used to interpret the collected images. These softwares are capable of making analysis to the collected images , and thus, obtaining vast amount of information. (Figure 1) (2, 4, 5)
Figure 1: Confocal microscope main components: pinhole, laser as light source, detector (photon multiplier tube) and beam splitter.(3)
Photon multiplier tube
Photon multiplier tubes convert photons to electric signals. They are characterized by high internal gain and they have sensitive detectors for the low intensity applications, for instance, the fluorescence spectroscopy. The photon multiplier tube is composed of a photocathode and dynodes located in the evacuated glass tube . When a photon with adequate energy hits the photocathode, it will release an electron due to the photoelectric effect. these electrons will be accelerated towards dynodes and additional electrons will be generated in every time they hits the dynodes. This will lead to more and more multiplication of photons as long as they getting in contact with the dynodes cascade series. This cascading effect creates 105Â to 107Â electrons for each electron (photoelectron) that is ejected from the photocathode. Amplification is dependent upon the number of dynodes as well as the accelerating voltage. This amplified electrical signal is collected at an anode at a measurable ground potential (Figure 2). (6) The photocathode is commonly made of mixture of alkali metals, which make the photon multiplier tube sensitive to photons all over the visible part of theÂ electromagnetic waves . The photocathode have a high negative voltage, ranging between -500 and -1500 volts.
Figure 2: Photon multiplier tube amplifies photons through hitting of electrons to the dynode series.(6)
Florescence and confocal microscopy uses
Confocal microscope is dependent on florescence and florescence probes ion acquiring images. florescence technology can be used in imaging various structures of the cells, for instance, mitochondria, cytoskeleton and filaments, endoplasmic reticulum, golgi apparatus, , and nucleus. Florescence probes are also used in studying dynamic processes and local environment variables, such as concentrations of inorganic metallic ions, studying pH, membrane potential , and reactive oxygen species (ROS). Confocal microscopy is also used in studying cellular integrity, apoptosis, endocytosis, exocytose, membrane fluidity, protein trafficking, signalÂ transduction, and enzyme activities. Moreover, fluorescence probing is used in molecular genetics, namely in genetic mapping and chromosome analysis. (6)
Considerations in sample preparation for confocal
Fixation: Fixation have to be adequate enough to stabilize the antigenic sites as well as other tissue components. If fixation is not done properly, florescence of the sample will diffuse and disappears over a very short time. Most commonly used fixative is buffered paraformaldhyde at 4oC. The paraformaldhyde can be either freshly prepared from powder or bought in vials sealed with inert gas. The commercial formalin is not recommended as it contain methanol as well as other impurities that can induce improper fixation. Methanol and acetone are not good fixators as they precipitate specimen proteins, resulting in shrunken and damaged tissue specimens.(7)
Immunolabelling : It is after fluorophores excitation. . Excitation and emission spectra of the used fluorophores must synchronize with the filters.
Slide preparation and mounting: The mountant you use under the coverslip is important for several reasons, namely, sensitivity of florescence to pH (absorption and emission are maximal at pH 8.5), preventing photo-bleaching by using additive to mountantÂ , and finally, the of the mountant refractive index has to be as close as possible to that of the fixed tissue (1.515). (7)
Working distance of the lens : it is defined as the space between the front element of the objective and the top of the coverslip. Working distance is dependent upon magnification and the objective numeric aperture. The working distance is in inverse relation with magnification. (7,8)
Advantages over conventional microscopes:
Through the concept of pinhole, confocal microscopes gained several advantages over the conventional optical microscopy, namely, ability to reduction of background information from the focal plan, controlling visualization of image depth, ability to collect several optical sections from the specimen, ability to use relatively thicker specimens, very high image quality, three and four dimensional images interpretation, and capability to image both fixed and non fixed images. Nowadays, confocal microscope technology became of great importance and proves to be one of the most imortant emerging advances in optical technology ever.(1, 2, 3)
Confocal microscope made a true revolution in cell imaging, especially live cell imaging. Its main idea is the use of pinhole technique to detect a high quality image for the examined specimen.
2- Pasi Kankaanpää, Turku Bioimaging (More Than You Can imagine) , 2011
5- (Nathan S. Claxton, Thomas J. Fellers, and Michael W. Davidson, LASER SCANNING CONFOCAL MICROSCOPY)
8- Müller, M., 2006. Introduction to Confocal Fluorescence Microscopy. SPIE Press.
Two Photon Microscope
Two photon florescence excitation
Two photon florescence excitation is defined as a florescence procedure in which a fluorophore is excited when simultaneous light absorption of two photons occurs (Figure 1). (1) In general, photon florescence occurs when exciting a fluorophore (A florescent molecule) from ground energy state to an excited higher energy level state with a photon. In conventional confocal microscope, single photon excitation occurs. The florescence occurs when the fluorophore absorb This single photon, which is is emitted from either two photon confocal microscope, two photon excitation occurs due to simultaneous absorption of two photons emitted from infrared spectral range. (2) An information to be mentioned here is that photons emitted from ultraviolet spectral range of normal confocal is known to be of higher energy than those of photons emitted from infrared spectral range in two photon. (1) The two photon excitation is not common to occur, accordingly, it only happens at high density photons exists (focal point) . The end result will be accurate excitation at focal plane. Neither above nor below it. This allow high degree of tissue penetration because excitation limited to focal plan provides less phototoxicity to the sample. According to the above, image of to photon microscope is characterized by the ability of optical sectioning as in normal confocal, and in addition, it provides with high degree of tissue penetration which is not accessible in normal confocal. (1,2)
Figure 1. Comparison between one photon and two photon excitation. in normal confocal only one photon is excited while in two photon there are 2 photons which lead to accurate excitation at focal plane.(1)
Linear and nonlinear optical microscopy
Linear and non linear excitations can be differentiated in optical microscopes. Linear excitation is the type of excitation found in conventional confocal microscope where single photon is used. On contrast, and in multiple photon microscopes, including the two photon, non linear excitations leads to verifying novel image properties.(3, 4)
Comparison between 2 photon microscopy and conventional one photon confocal microscopy
Two photon microscopy has many advantages in comparison to conventional one photon microscopy. A major advantage is the high penetration ability of two photon microscope which allows the obtaining of optical sectioning from deeper tissues. This advantage is very important in intravital florescence imaging. It is assumed that two photon improves image penetration by at least 2 fold more than the normal one photon confocal microscope. Another advantage of two photon over the normal confocal is reduction of overall photo-bleaching and photo-damage due to limiting it to narrow region around the focal plane. On the contrary, all the focal planes in the one photon confocal microscope are exposed to the excitation light in every time it perform optical plan image collection. In other words, it could be mentioned that normal confocal exposes the tissue to risk of photo-bleaching and photo-damage in every optical plan collection. This make two photon microscope more convenient when collecting three dimensional data (Z series) that requires many optical plan scans. A third advantage isn on using endogenous (for example, nicotine adenine dinucleotide) and exogenous (for example, ion-sensitive indicators) fluorophobes that requires ultraviolet excitation. On using two photon microscope , NADH and ion concentration in single cells and intact tissues are successfully measured without much risk of photo-bleaching and photo-damage, which is not the cases in normal confocal. Finally, two photon microscope has shown great success in three dimensional florescence imaging for specimen, three dimensional localized photochemistry at a subfemtoliter scale, such as photo-activated release of caged calcium ionsÂ or neurotransmitters, Â and measurements of three-dimensional mobility of fluorescent molecules by using the fluorescence photo-bleaching recovery. (5,6)
Uses of two photon microscopy
Two photon microscope can be the best choice when it is needed to visualize the cells and cellular interactions in their in vivo native environment. Additionally, It is of great benefit when tissue fixation and labeling is not applicable, foe example in signal loss or undesirable changes of the sample. Despite most fluoresce proteins can make two photon excitation, two photon microscopy can be used in tissues with non florescent proteins through second and third harmonic generation phenomena that enable detecting specific tissue structures without the need of labeling. Two photon microscopes has been widely used in biological studies. It is very practical and efficient in experiments studying cell migration, cells and immune interactions between cells in infections, neuronal activity, either through brain slices or intra-vital imaging and it is also used in studying cellular signaling for vital tissues.(2)
Promising Applications of two photon microscopy in biology
Two-photon microscopy is expected to have an impact in various fields of science such as physiology, neurobiology, embryology and tissue engineering, in which the imaging of highly scattering tissue is needed. Thanks to the two photon microscope, opaque tissues such as human skin can be detected with high degree of cellular detail . In clinical practice, two-photon microscopy can be applied in noninvasive optical biopsy that requires high-speed imaging procedures. This can be achieved using video rate two-photon microscopy . Two photon microscopes applications can be very promising In the field of cellular biology, particularity in using two-photon excitation to produce localized chemical reactions, such as in three dimensional resolved un-caging and photo-bleaching recovery studies.(1)
Two photon microscopy is a confocal microscopy variant with much abilities in in-vivo cellular imaging applications due to the technique of 2 photon excitation instead of one.
1- Peter TC So , Two-photon Fluorescence Light Microscopy
2- Tibor Veres, Turku Bioimaging (More Than You Can imagine) , 2011
3- Helmchen, F., Denk, W., 2005. Deep tissue two-photon microscopy. Nat. Methods 2, 932-940.
4- Bush, P.G., Wokosin, D.L., Hall, A.C., 2007. Two-versus one photon excitation laser scanning microscopy: Critical importance of excitation wavelength. Front Biosci 12, 2646-2657.
5- Rubart, M., 2004. Two-Photon Microscopy of Cells and Tissue. Circulation Research 95, 1154-1166.
Förster Resonance Energy Transfer (FRET)
Introduction about FRET
Förster or (Fluorescence) Resonance Energy Transfer (FRET) is a physical phenomenon that was discovered over 50 years ago. Its use is biomedical and drug research is increasing over time. FRET is distant dependent phenomenon based on the transfer of energy from a donor molecule to an acceptor molecule. It is used to detect molecular interaction depending on of the fact that it is sensitive to distance. In FRET technology, donor molecule is the used dye, while the acceptor is the chromophore. Energy is absorbed by the donor and transmitted to the acceptor. This interaction occurs in distance greater than the interatomic distances, and there is no either change to thermal energy or molecular collision. This transfer of energy results in a reduction in the donor's fluorescence intensity and excited state lifetime. On the other side, it leads to increasing the emission intensity of the acceptor. The pair of molecules that interact in FRET is called donor-acceptor pair. 1
Förster resonance energy transfer (FRET) has shown great progress in detecting proteins dynamic interactions. The use of green fluorescent protein (GFP) targetingÂ in FRET mechanisms has been widely used achieving notable success. FRET between two differently colored fluorescent molecules has been vastly used as well . 2, 3
Concept and Mechanism of FRET
FRET phenomenon happens when the separating distance between two fluorophores is less than 10 nanometer (nm).2 It depends on a mechanism in which a donor fluorophore in an excitation state, transfers its energy to a near acceptor chromophore in the form of non-radiative energy transfer through a dipole to dipole interaction. The theory of energy transfer depends on the idea that treating an oscillating dipole, which is capable of energy exchange, with a second dipole of a similar resonance frequency.4
In FRET, acceptor is not necessarily to be florescent, yet it is often so. If the donor dye is florescent and acceptor is not, the measurable parameter is the increase of donor's fluorescence. Conversely, and if both donor and acceptor are both fluorescence, the measurable parameter is the ratio of change in fluorescence intensity between donor and acceptor.5
In other words, FRET can be detected via either measuring florescence time of donor/acceptor fluorophores, or intensity change.6 The lifetime of florescence is defined as the time spent by a fluorophore in its excited state.5
On excitation, electron will jump from its ground state (called S0) to a higher oscillation level. Afterwards, they will drop rapidly (in picoseconds) to the lowest oscillation level (S1) and then, returns back to the original (S0) state in nanoseconds time emitting a photon of light. This photon will be absorbed by the donor molecule, and afterwards, a radiation-less energy transfer will happen between this donor molecule and the acceptor molecule nearby. (Figure 1). 1
Figure1: FRET : A radiation-less energy transfer occurs between donor and acceptor.1
FLIM (Fluorescence lifetime imaging microscopy)
FLIM is an accurate way to measure FRET. An advantage for FLIM is that crosstalk artifact are less likely to happen. The basic concept of FLIM is dependent on acceptor photobleaching. The fluorescence of the donor dye is quenched by FRET, and after, quenching amount can be estimated by measuring the decrease of the fluorescence decay time of the donor in the presence of FRET. Accordingly, it can be said that FILM is a sort of efficient FRET.2 FLIM is sensitive, and give convenient interpretation to changes in specimen, like polarity and pH. 5 However, there are several limitations preventing FLIM to be always dominant over FRET. One thing, is the difficulty in measuring nanosecond lifetimes due to expensive instruments used, their unavailability, and the difficulties to maintain them. Other limiting disadvantages of FLIM include speed; FILM is known to be slow, and it takes several minutes in order to acquire a single image. However, these disadvantages might change in the future with developing more user-friendly, cheaper and faster systems. 2
Use of FRET in biology
FRET microscopy applications in live cell imaging are much fruitful and they are expected to expand in the following years. To date, many studies were performed using FRET technique and achieved great success. Using florescence proteins as FRET donor and acceptor has shown a notable importance as it can be used in live cell imaging. For example, detecting dynamic processes in membrane while they happening as well as facilitating visualization of proteins inside the viable cells. FRET is used to detect various protein interactions between two fluorescent dyes labeled proteins, such as visualizing oligomerization of receptorsÂ Â and transcription factor interactions. These procedures are hard to analyze in the biosensor work due to complicated nature of proteins composition . However, these experiments can be informative with suitable experimental manipulations. Another example to mention is overcoming problem of identifying membrane micro-domains, in which resolution limit of conventional confocal microscope is unable to interpret. Additionally, FRET is used in assays of intracellular Ca2+, cAMP activityÂ Â and protease activityÂ . 2,7
FRET is a technology depending on radiation-less energy transfer between donor and acceptor molecules. It a is efficiently used in dynamic imaging and detecting various interactions inside the viable cells, and future is promising for more advantageous applications of FRET.
2- Piston, D.W., Kremers, G.-J., 2007. Fluorescent protein FRET: the good, the bad and the ugly. Trends in Biochemical Sciences 32, 407-414.
3-Karpova, T.S., Baumann, C.T., He, L., Wu, X., Grammer, A., Lipsky, P., Hager, G.L., McNally, J.G., 2003. Fluorescence resonance energy transfer from cyan to yellow fluorescent protein detected by acceptor photobleaching using confocal microscopy and a single laser. Journal of Microscopy 209, 56-70.
6- Turku Bioimaging (More than you can imagine), 2011
7- Loura, L.M.S., Fernandes, F., Prieto, M., 2009. Membrane microheterogeneity: Förster resonance energy transfer characterization of lateral membrane domains. European Biophysics Journal 39, 589-607.
Spinning disk is a novel and a rapidly growing technology used mainly in dynamic live cell imaging, yet it can also be used in fixed cells. This techniques allows the study of intracellular dynamics through a three dimensional high quality imaging, with high speed, and accordingly, lesser exposure to light that result in photosensitivity and photobleaching.1,2
Concept of spinning disk microscope
Spinning disk microscope's main is the presence of disk (spinning disk) that contain thousand of minute pinholes in it. These disks are rotating ones with holes arranged in spiral manner. Actually, there are 2 disk with a dichrotic mirror in between. The first disc is called the pinhole disk (Nipkow disk), while the second is called lens disk and it contains thousands of micro lenses arrays in order to increase sample illumination. From the above, it can be stated that spinning disc is a kind of confocal microscope having thousands of small, simultaneously rotating pinholes instead of a single pinhole in the conventional confocal. These pinhole array is capable is of collecting fast, multiple, and spontaneous images in the same time it takes to collect a single image by using normal confocal. Accordingly, much desirable results can be obtained in dynamic cell images. CCD camera is important component in this imaging device with it role in collecting and interpreting and the pinhole images. The higher the sensitivity of the CCD camera, the faster the ability to collect faster frame rates, and accordingly, the faster the spinning of the discs. 2,3, 4
Design and action of spinning disk microscope
In the microscope, there is the Nipkow disk (figure 1), which contain several pinholes and parallel slits that are arranges in a spiral manner. The illumination of pinholes and the slits are from the back of the disk in the side opposite to the eye. Florescence light beam emerging from the sample will pass through the objective lense in its way to the spinning disk pinholes. When the Nipkow disk rotates, the sample is scanned by the thousands of pinholes arrays. The light comes out from illuminated points in the sample is focused by the opposite disk through the microarray of lenses. Afterwards, light will pass through the dichrotc lens (dichrotic beam splitter) and passes through barrier filters on its way to the monochrome CCD camera. The entire specimen will be scanned in the same way producing a three dimensional dynamic interpretation.4
Spinning disk technology is very efficient. Each Nipkow disc contains average of 20 000 tiny pinholes with a spinning speed up to 5000 rotations every minute. In every millisecond, each excited light beam column passes through 1000 tiny pinholes. This huge speed can create movies with speed of 2000 frames per second (fps), and thus, allowing high level of detecting real-time events in the cell. 5
Figure 1: Spinning disc (Nipkow disk) containing thousands of pinholes arranged in spiral manner. 4
Some advantages of spinning disc over laser confocal
In normal laser confocal microcopies, the most common configuration for the scanning is depending on using a pair of oscillating mirrors driven from galvanometer so that to maintain sustained deflection of the laser beam. This process is known as scanning. On the other side, and in spinning disc microscope, one of the mirrors lead the excitation and emission light along the fast access (horizontal in direction), and the other mirror direct light towards the slow axis (vertical in direction). Regarding to scanning speed, spinning disc is much faster than the confocal. Another important difference is that scanning disk microscopy is much practical in scanning slower events that happen in many seconds. The spinning disk forms the image in sequence of single pixel per time, while the laser confocal form image by spontaneous illumination of the whole specimen through parallel pinholes. Partial rotation of Nipkow disk in the spinning disk microscope can scan the sample by almost one thousand light beams which are capable to scan the whole image plane in less than one second with high degree of accuracy in comparison to the normal pinhole confocal. (Figure 2) 4
Figure2: Laser confocal microscope (a) and spinning disc confocal microscope (b) 4
Use of spinning disk
Spinning disk is the optimal technique to obtain images for rapidly occurring cellular processes in dynamic live cell imaging with minimal photo-bleaching and photosensitivity.2
But what are disadvantages of spinning disc microscopy?
Spinning disc has some disadvantage that can limit it's usage, namely, the pinhole crosstalk artifact which increase the signal coming from background of thick specimens. and consequently reduces the axial resolution. Another disadvantage is the low light transmission level through the Nipkows disc. This makes it not much convenient in less illuminated florescent samples. Additionally, low field uniformity can another problem which requires correction lens or liquid light guide to homogenize the arc source. Finally, lack of ability of detecting field of interest in the high illumination power after low intensity imaging. This limits it's usage in photo-bleaching and photo-activation. 4
2- Turku Bioimaging (More than you can imagine), 2011
Contrast microscopes has many variants such as phase contrast microscope, dark-field microscope, bright-field microscope and differential interference. The most important variant is phase contrast microscopy which was invented to the first time in the year 1934 by physicist Frits Zernike from Deutschland. Frits obtained noble prize for his invention in the year 1953.(1, 2) Contrast microscopy is an optical technique depending on contrast-enhancing and can be used in transparent specimen to produce high contrast images. Phase contrast microscopy works by mean of effecting optical path of light to enhance contrast of the transparent and colorless objects in order to be more visible. This enables to visualize components of cell and bacteria which are hard to see by conventional optical procedures.(2) Other examples can be cell in cultures, microorganisms, thin tissue slices, lithographic patterns, fibers, latex dispersions, glass fragments, and sub-cellular particles including the nucleus and other cellular organelles).(1)
Concept of work
It is known that high refractive index structures bend light with a greater angle than low refractive index ones. It was also noticed that light is delayed by more or less a quarter wavelengths during its refraction passage. On applying this to light microscope with bright field mode, we will find that highly refractive index structures refract light away from the center of the lens to the periphery in comparison to those with low refractive index ones with an approximate delay estimated by a quarter wavelength. Accordingly, It can be said that light arising from most of structures passes through both center and periphery of the lens according to their refractive index profile. On these bases, it can be assumed that if light from an object has arrived to the edge of lens has been delayed by half wavelength from the light arrives to the center of lens, so the light rays are said to be out of phase by a half wavelength. Both light waves will cancel each other when objective lens shows the image in the focus. Also, a reduction of object's brightness will be observed. Degree of brightness decrease is dependent on refractive index of the object.(3) Phase contrast microscopy is not sensitive to polarization and birefringence effects, which is an important advantage on doing examination to living cells grown in plastic tissue culture vessels.(4)
Condenser annulus and phase plate are pivotal components in phase contrast microscope
Condense annulus is a special annular diaphragm, having similar diameter to the internal phase plate and it is optically conjugated to it. It is located in the front focal plane of the condenser. Condense annulus is made from an opaque, flat and black plate which absorbs light, and a transparent annular ring found also in the front focal plane. The sample can be illuminated by a non-focusing, parallel light coming out from the ring. The condenser annulus either replaces or settles close to the iris diaphragm located in the front aperture of the condenser lens. On making phase contrast experiments controlling the condenser with annulus and aperture diaphragm, it has to be checked that the iris diaphragm is wider than the periphery of the phase annulus. The amount and location of the detracted light is proportion to the number, size, and refractive index of the light scattering components in the specimen. For majority of specimens, only a small part of the incident light waves are diffracted, while the majority of the light travels with no deviation to illuminate the whole plane of the image. Another component is theÂ phase plate, whichÂ is found in or near the objective backward focal plane in order to selectively change the phase and amplitude of the surrounding light passing through the specimen (Figure 2).(4)
Figure 2: Controlling of condenser annulus and phase plate to light beam, amount of light and image formation.(4)
Examples for using phase contrast microscopy
Phase contrast is better than bright field microscopy in higher magnifications. Additionally, it is more convenient when and the specimen is colorless or when details are very fine so that the colors of the sample do not clearly appear. Cilia and flagella, for example, are nearly invisible in bright field but are satisfactory visible in sharp contrast and in phase contrast. Amoebae appear as unclear outlines in bright field, and conversely, it appears very detailed on using the phase contrast microscopy. Most living microscopic organisms are much more clear to visualize in in phase contrast. (3)
Another example could be the cells in human glial brain tissue. These cells grown in monolayer culture an put in a nutrient medium containing amino acids, vitamins, mineral salts, and fetal calf serum. In the conventional bright field illumination, the cells appear as half transparent cells. Only the high refractive regions in these cells are visible, for example membrane, nucleus, and unattached cells. Conversely, and on viewing these cells using phase contrast optical accessories, the same field of view shows significantly more and more details to the containing structures. Cellular attachments become easy to detect, as well as much of the inside structures. In addition, the contrast range is obviously improved (Figure 1). (4)
Figure 1: showing cells by normal bright illumination (a) and by phase contrast microscopy (b). (4)
Differential Interference Contrast (DIC)
Differential Interference Contrast is another method of showing contrast in the unstained specimen. The advantage of differential interference contrast is obtaining a brightly appeared object against dark background with avoiding halo diffractions that happens with normal contrast microscope .(5)
Dark field microscopy
Dark field microscopy is another variant for contrast microscopes, and it can be used as cheap alternative as well. Operating expenses of dark field are much lower than that of phase contrast as of optics of dark field are much cheaper than that of phase contrast, yet a high quality with high resolution image can be still obtained.(6)
Contrast microscopes are optimal in viewing of colorless and transparent specimens, as it gives highly distinguishable, definite and distinct image for such specimens.
Atomic Force Microscopy (AFM)
In the year 1982, scanning tunneling microscope (STM) was invented by Gerd Binning and Heinrich Rohrer at IBM Zurich. STM is considered to be the ancestor of all scanning probe microscopy techniques. After 5 years, they got Noble prize for this invention.(1) After 4 years of inventing STM, and in the year 1986, Binnig, Quate, and Gerber invented the Atomic Force Microscope.(2,3) The Atomic Force Microscope was invented to solve with an important problem in scanning tunneling microscope (STM) - that it only interpret images for conducting or semiconducting surfaces . (2) The main component of AFM is called cantilever. In which its free end contains a tip that scans the surface of the required sample. The forces between the fine tip and the sample are detected by AFM, and accordingly, image is interpreted. ATM has the advantage of imaging nearly all surfaces types, such as polymers, ceramics, composites, glass, and biological samples. (2)
Concept of atomic force microscopy (AFM)
In atomic force microscopy (AFM), a probe scans the sample in order to get information about the sample's surface. It is a kind of mechanical microscope with idea of operation resembling the old record turntable, by which there is a sharp tip that is attached to a cantilever with spring. This cantilever senses the force between the tip and the specimen.(4) The information collected from the surface-probe interaction can be either simple information; such as physical topography, or variable and complex; such as measuring physical, magnetic and chemical properties. The data detected by the probe is scanned as dotted (raster) information, and interpreted in the form of map. The AFM probe has a very sharp tip which is usually less than 100 Angstroms in diameter. This tip is found at the end of a small cantilever. The cantilever is attached to a piezoelectric tube that controls scan movement all over the sample surface. While scanning, cantilever deflects according to surface topography of the sample due to interaction forces between tip and sample surface. Defection is measured by laser, then information is interpreted via computer, which generates a map of topography and other needed properties for the sample . Large areas (up to 100 Î¼m square) as well as small areas ( less than 100 nm square ) can be interpreted. Components of AFM can be seen in Figure 1.(5)
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Figure 1: AFM and mechanism of image acquiring through: scanning surface cantilever, piezoelectric tube and laser.5
Modes of Action in AFM
AFM has many modes of action. For example:
Contact Mode:Â It is the first and the most common mode In AFM. In contact mode, the tip scans the surface and it is deflected on moving over a surface corrugation. In constant force mode, the tip is constantly adjusted for constant deflection with a constant height from the surface, and then, displayed as data. Nevertheless, feedback circuit can imitates the ability to scan the surface in this manner. This method is known as variable deflection mode and it can be used in small, high speed atomic resolution scans.
Lateral Force Microscopy: It measures frictional forces applied on a surface. By measuring the cantilever's twist, and not just its deflection. This method can detect areas of higher and lower friction in qualitative manner.
Noncontact mode: It is one of the AC modes that uses an oscillating cantilever. A stiff cantilever is oscillated with its tip is very close to the sample, but not touching it, and this is why it is called non-contacting. The forces between the tip and the sample are low. The concept of estimation here is measuring changes happens to resonant frequency or the amplitude of cantilever.
Tapping mode (Also called Intermittent contact or Dynamic Force mode (DFM) ) AFM: This mode has many names, in which the most common name is tapping mode and others are dynamic force mode or intermittent contact mode. A stiff cantilever is oscillated closer to the sample than in noncontact mode. In this mode, the tip of the cantilever taps the surface in an intermittent manner. Stiff cantilevers have to be used to avoid the stuck within the water contamination layer. Advantage of tapping is to ameliorate lateral resolution on soft samples.
Force Modulation: This mode detects the material properties through interaction between the sample and the tip, though which the tip oscillates at a high frequency and inter in a repulsive state . Sample elasticity is calculated by measuring The slope of the force-distance curve. The data can be interpreted together with topography, which compare both the height and the properties of material.
Phase Imaging : In this mode, the phase shift of the oscillating cantilever relative to the signal detected. The resulted phase shift can be correlated with specific properties of the material that influence the tip/sample interaction. The phase shift can be used to detect different areas in the sample having different properties, such as friction, adhesion, and viscoelasticity. It can be used together with the dinamic force mode to measure topography as well.(2)
Uses, Advantages and disadvantages
AFM can be used in various applications of nanoscience and nanotechnology. The advantage of using STM and AFM is their efficiency with samples in very high vacuum environment, as well as in ambient conditions and liquids. It is very practical to use AFM in biological systems as it can image non-conducting surfaces. The most important application of AFM is topography studies of and obtaining very high quality topographic image of the sample and in getting three dimensional surface images. It is needed when imaging is needed to be acquired under the natural condition of the sample. It can be used also on failure of optical microscopy to get the desired image due to inadequate resolution of these optical devices used. AFM is additionally capable of measuring nanometer scale images, and finally, it can be used in detecting variations in material properties such as surface stiffness, elasticity and adhesion. A disadvantage is decreased ability to study intracellular interactions and mechanisms.(1,4)
AFM can be perfectly used in scanning of surfaces with great accuracy.
3- Binnig, Quate, Gerber, 1986. Atomic force microscope. Phys. Rev. Lett. 56, 930-933.
4. Pasi Kankaanpää, Atomic Force Microscopy, Turku Biomiaging (More Than You Can Imaging), 2011
Fluorescence Recovery After Photo-bleaching (FRAP)Â is a method of determining the kinetics of diffusion in living cells by fluorescence microscopy. The method for this is done through following steps. The first step is to label the specific cell component required to be studied with a fluorescent molecule. Second, imaging of that cell is done. Third, a small portion of the cell has to be photo-bleached, and then detect fluorescence recovery over time by taking several images. There is recovery of florescence as a result of the diffusion and active movements of molecules within the cell replace the bleached fluorophore with the unbleached molecules which were found in different part of the cell. Fluorescence in the bleached region will recover over time. From the previous, we can notice that two items are detected in FRAP imaging when the fluorescence returns to the photo-bleached area, First is amount of light returned in relation to amount of light before photo-bleaching and second is speed of the florescent molecules returning back (Figure 1).(1, 2)
In FRAP plotting graph, recovery time is plotted as x value, while intensity of signal in the area scanned in microscopy is plotted as y. The time of recovery is a parameter to detect the mobility of florescence molecules.(1, 2)
Figure 1: FRAP graph: X axis is time and Y is intensity.(2)
One of the used fluorescent protein in FRAP is green florescence protein (GFP). FRAP is and old technique discovered more than thirty years ago. However, the recent renaissance in FRAP is due to the appearance of GFPs, GFP variants and other fluorescent proteins that enabled tagging proteins of interest with minimum damage to cells that can happen when doing microinjection of fluorescent dyes.(3) The fluorescent dyes produce light with one wave length, after they have absorbed light of another wave length for example emitting green after blue wave absorption. Nevertheless, if the dye is exposed to high intensity blue light, the dye will "photo-bleach" meaning that the high intensity light has rendered the dye unable to fluoresce. This phenomenon leads to FRAP. The rationale behind this is using FRAP to measure the capability of the molecule to turn around over time. To do this, a fluorophore have to be attached covalently to the molecule in the sample , such as protein, lipid, carbohydrate.(5)
Making FRAP by confocal
Performing FRAP requires using custom built systems to make the required measurements. Applying FRAP in the laser-scanning confocal microscope has made this FRAP technique to be widely available. Images on the confocal microscope are obtained through scanning with a focused laser beam across the sample then recording the emitted fluorescence through a pinhole located to the front of the light detector. A method to photo-bleach using this system is to define a region-of-interest (ROI) at the highest possible zooming, then setting the laser power to its maximum with setting the laser attenuation power to zero level. The high zoom increases the dwell time of the laser on the bleached region per line scan (laser intensity increases proportionally to the square of the zoom factor), which therefore greatly increases the radiation per area. But a more advanced method is to use an acousto-optical tunable filter (AOTF) which can be found in more recent confocal microscopes, which allows rapid (microsecond to millisecond) attenuation of the laser as it scans a field. AOTF allows accurate measurements of diffusion rates in defined areas through allowing rapid switching between the bleaching and normal beam.(3)
Application of FRAP
FRAP is used for imaging and measuring rates of intracellular molecular dynamics. It has been used, for example, to image microtubule and cytoskeletal dynamics and the mobility of membrane proteins.(6) FRAP can be used in detecting diffusion of molecules, visualizing active movement of cell components, recycling of the cellular components, studying movement behavior of fluorescence labeled molecules, for example, if these molecules move freely or bound to another static object.(1)
Fluorescence Loss In Photo-bleaching (FLIP)
FLIP (Fluorescence Loss In Photo-bleaching) is a technique very similar to FRAP, and it is used to to measure molecular mobility and dynamics in living cells. A certain known area is repetitively bleached over time by a high intensity laser beam and the area around it is checked for a regression in the level of fluorescence. This is applied instead of monitoring the recovery of fluorescence in FRAP. Any part of the cell connected to the area of bleaching will be gradually fading out due to the movement of bleached molecules towards that region. On the contrary, the cell parts which are nor connected to the bleaching area will continue to florescence and will not be affected. The FLIP technique can be used to assess if a tagged bio-molecule moves to a certain part of the cell. PA-GFP (Photo-activated green fluorescence proteins) are often used as an alternative to FLIP to see the dynamic movement of a bio-molecules in the cell. The negative part here is that tagging protein of interest with PA-GFP requires giving re-cloning steps.(3, 4)
FRAP is a technique based on detecting recovery after photobleaching in specimens with florescence molecules, and accordingly, it is used in experiments involved with cellular dynamic movements.
3- (NPG web focus: imaging in cell biologyâ€¯: Review [WWW Document], 2012. . URL http://www.nature.com/focus/cellbioimaging/content/full/ncb1032.html)
4- Rachel Underwood, Nikon note.
Stimulated Emission DepletionÂ Microscopy (STED)
STED stands for stimulated emission depletion microscopy. Its importance comes from that it was the first to emphasize that diffraction barrier could be broken via the florescence microscopy. STED was invented in the year 1994 by the scientist Stefan Hall in Turku, Finland (1). It is one of the principles of RESOLFT (reversible saturable optical (fluorescence) transitions), which is the utilization of reversible saturated optical transitions (fluorescence). These principles states that is florescence marker can change between two states; A and B states, these states are clearly noticeable and changeable, in which there is one florescence state, while the A and B transitions are light driven.(1,2)
Concept of working in STED
Florescent markers in STED microscopy are excited by a laser, typically like in the conventional laser scanning microscopy. This laser is scanned across the examined sample. The temporarily switching off to the marker's florescence molecules located in the outer area away from focus will result in formation of small florescence area. This can happen by second, red-shifted STED beam which use stimulated emission to let the molecules in their dark ground state. The area in which fluoropobes are not switched off decreases regarding to the size of the molecule, when there is an increase in the intensity of STED beam. Conversely, on blocking STED beam, it will operate as standard conventional confocal microscopy.(3) Figure 1 shows the image obtained from STED versus normal laser confocal microscopy.(4)
Figure 1: STED (b and d) vs Confocal (a and c) 4
In the STED microscopy, fluorophore will absorb the photon so that