I added a new page named “Terms list“.
It is a list of terms (duh!) with a brief description.
If you have any comments and suggestions, please comment here.
I added a new page named “Terms list“.
It is a list of terms (duh!) with a brief description.
If you have any comments and suggestions, please comment here.
During training on the FACS machine in our faculty facility, I encountered the Fluorescence Spectrum Viewer from BD bioscience.
It is a JAVA interactive site that allows to to view excitation/emission spectra of up to 8 differnt fluorophores simultaneousely.
It lets the user to choose the desired excitation laser, and shows the excitation & emission spectra *to that laser*. moving the cursor over the spectra gives you actual numbers of % excitation & % emission efficiency.
You can use filters to test which filter is most suitable for your fluorophores.
This site is intended for FACS users, so you can choose from any one of a number of cytometers from BD. However, I think this site can be usefull for microscopists too.
EDIT: I found more sites like that, each with its own set of fluorephores –
Fluorescence SpectraViewer at Invitrogen (Life technologies) site.
Fluorescent Dye Spectra at U Arizona (only chemical fluorophores)
Evrogen Spectra Viewer at Evrogen (only a very limited list of fluorescent proteins)
There is also this Cell staining tool from invitrogen. The concept is cool, but I think it is a very limited tool and I don’t think it is very helpful.
Nowadays, confocal microscopy is possibly the most widely used optical method in biological research. This methods creates better (and prettier) images than widefield microscopy (whether transmitted light or epifluorescence). The main advantages of confocal vs. widefield microscopy is the elimination of out-of-focus glare (thus increasing resolution and increasing signal-to-noise ratio) and the ability to collect serial optical sections of the specimen (z-sections).
The basic configuration of the optics is similar to that of the epifluorescnece microscope. The addition that created the confocal microscope, invented by Marvin Minsky in 1955, was to add two pinholes. The light produced by lamp (or laser) passes through the first pinhole on the way to the specimen. The light that is reflected (bright light) or emitted (fluorescent light) from the specimen passes through a second pinhole on the way to the detector (eyepiece, camera or any recording device). The two pinholes have the same focus – thus they are confocal. The light from other focal planes cannot go through the second pinhole, and this reduces the background “glare” of out-of-focus fluorescence seen in epifluorescence widefield microscopes.
Since biological samples usually have thickness of a few microns at least, one can get an image of a thin slice of the sample (e.g. 0.1 µm) without physically slicing the sample (optical section). We can then move the focus along the Z axis to get clear images of up or down sections. Thus, for a cell 3µm thick, we can have 30 hi-resolution images 0.1 µm thick from bottom to top (z-sections). These images can then be stacked one on top of the other (z-stacking) to create a single 2D image or to reconstruct a 3D image of the sample.
Here’s an example from an experiment I did last week (note that this is a widefield, not confocal microscope):
Above is a composite image of 31 Z sections of U2OS cells, create by the ImageJ program. The”pseudo-blue” represents the blue fluorescnce of a dye called DAPI (4′,6-diamidino-2-phenylindole) which intercalates into DNA, and is therefore a popular nuclear dye. When bound to DNA, it is excited by UV light (peak at 358nm) and emits blue/cyan light (peak at 461nm). The “pseudo-red” color represents the fluorescence of mCherry-ZBP1 fusion protein. mCherry is an RFP.
You can see in the image that the first and last few images in the series are out of focus. You can therefore choose the best or sharpest Z-section according to your needs.
However, most people do not show a single Z section since then we miss a lot of information that is found in other sections. The available programs today allow “stacking” the section to create a projection of all the sections into a single image.
Here is the maximum projection of the Z sections shown above:
Maximum projection means that the algorithm chooses, for each pixel, the highest value found in any of the 31 Z sections. However, since we chose all 31 sections, we can still see a “glare” or halo. This is a result of the “halos” from the out of focus sections.
I therefore choose only a few sections to create the next image:
This image is now sharper and better looking.
The program allows you other options besides maximu projection: you can choose minimum, average, median, and even standrad deviation, seen in the next image (DAPI channel only):
It looks very cool. I stacked the entire 31 sections (of a differnt field), so you can see the halo from the out-of-focus sections sorounding the “black” rim of the nucleus (black since it has the minimal standrd deviation value for all the images). The blue zones, with high SD, suggest a larger differnce in fluorescence between the differnt sections.
Above, I mentions that the blue and red are pseudo colors. What actually happened was that the images I took with the microscope at each channel (range of wavelengths) is actually maintained as a greyscale image.
Using the image analysis program you can then merge the images of the differnt channels (up to 4 in ImageJ) to create a color image. When creating the merged image, you determine what color to assign to each channel. Here is the same image, but with the colors reversed:
You should take that into account when you see pretty pictures in sceintific journals.
The program also allows to creade 3D representations of your Z stack. but I haven’t learned how to do that.
There are many other tools that one can use with the image analysis program besides creating the image. One important feture is the ability to measure the intensity of the fluorescent signal (actually, the pixels) in certain areas within the cell. You can measure distances and angles between objects and probably many moer that I still have to learn.
While reading about microscopy, I came across mentions of a parameter called numerical Aperture (NA).
NA is a measure of the ability of the objective lens to gather light and resolve it, at a fixed distance from the specimen, through a specific media (e.g. air, water, immersion oil).
As shown in the figure, light from the specimen enter the objective in an inverted cone as seen in this figure:
The angle µ is half the angular aperture A. NA is calculated by the following equation:
NA = n(sin µ)
n is the value of the refractive index of the medium between the specimen (the cover glass) and the objective. The value of n is between 1.0 (for air) up to 1.51 (for special immersion oil).
When imaging through air (usually with low magnifications), then n=1 and so NA depends solely on the sin of µ which theoretically can be a maximum of 1 (sin(90°)=1). In practice, it is difficult to achieve NA>0.95 with “dry” microscopy (i.e. through air).
Therefore, many microscope objectives (mostly for high magnifications) are designed to be used with water (n=1.33), glycerin (n=1.47) or immersion oil (n=1.51).
So how does the NA relate to resolution?
The resolution of an optical microscope is defined as the shortest distance between two points that can still be distinguished by the observer. This can be perceived by the concept of Airy diffraction patterns (named after physicist George Biddell Airy).
I will not go into the physics of it now (since I need to learn it first). It is enough to say that point sources of light in the specimen (e.g. fluorescent proteins) appear as bright disks, surrounded by bright concentric rings.
The limit of resolution is determined by the ability to distinguish between two closely spaced Airy disks. The 3-dimentional representation of the Airy disks is called the point spread function. The resolution of the microscope is determined (in part) by the NA of the objective. You can explore it yourself using the Java tutorial of NA & resolution at MicroscopyU. You can see that the larger the NA, the better the resolution.
The resolution is determined not only by the NA of the objective, but also of the sub-stage condenser. The alignment of the two, to create an accurate light cone, is also important.
Another parameter that greatly affects the resolution is the wavelength (λ) of light that is used. Generally, the shorter the wavelength, the better the resolution. There are three equations developed to describe the resolution (r):
r = 0.5λ/NA
r = 0.61λ/NA
r = 1.22λ/(NA(obj)+NA(cond))
NA in the first two equations refers to the general NA of the microscope.
These equations were calculated based on multiple optical factors and, in some instances, the actual resolution may be even better than the limit calculated by the equations.
However, other factors, like background, contrast, illumination and more can also affect the resolution.
The practical limit (for green light) is ~250nm.
Just to put things in perspective, the size of an epithelial cells is ~30µm, that of a yeast cell is ~4-5µm. The size of the yeast nucleus is ~1µm. A chromosomally packed DNA length of 100kb (100,000 bases) is ~500nm, when stretched. Viruses are usually ~30-50nm. A single ribosome is also ~30nm. A single average protein will be ~5nm in size. Therefore, viewing single molecules in cells is a real challenge.
Now that we know a little bit about fluorescent proteins, let us talk about application. The first and most obvious application is fluorescent microscopy. We will discuss other applications in future entries.
The basic function of a fluorescent microscope is to shine the sample with a specific, desired, bandwidth of wavelengths and then detect (and record) the much weaker emitted light of the excited molecules.
If the microscope is properly designed, only the emitted light is detected by the eye or the detector, superimposed on the dark background of the specimen. The limit of detection is usually determined by the signal to noise ratio, i.e. the darkness of the background compared to the emitted light, and the exciting light (which is usually 5-6 orders of magnitude stronger than the emission).
The simplest design is the epi-fluorescence microscope, which consists of a light microscope in which light reflected from the sample is at a longer wave-length than that of the excitation light. The basic principle is of a vertical illuminator in which one side contains the light source (called epi-fluorescence or episcopic lamphouse) and the filter cube turret at the other end (between the objective and the eyepiece/camera). Multispectral light, usually from an arc-discharge lamp, passes through selective excitation filter (usually ultraviolet, blue or green) to produce a bandwidth of the desired wavelengths. The light is then reflected from a dichromatic (often called dichroic) mirror or beamsplitter, through the microscope’s objective to illuminate the sample with intense, specific, light.
If the sample fluoresces, the emission light that is gathered by the objective passes back through the dichroic mirror. The beam is then filtered again by the emission filter (or barrier filter) which blocks the unwanted excitation light that is reflected by the sample.
One advantage of the epi-fluorescent microscope is that the excitation light passes through the objective lens (which, in this case, acts as a condenser), and the emission light goes through the same lens back to the eyepiece/detector. Because the same, single part of the microscope is used both as condenser and as objective to view the sample, there is always perfect alignment.
Another advantage of this microscope is that it is cheap (relative to other, more advanced microscopes). It also requires low maintenance, low power, takes less space and relatively easy to operate. Thus, this microscope is suitable for labs with low budget, and no need for high-resolution, low background, pictures.
However, the epi microscope’s major drawback is the background fluorescence. Since the excitation light passes through the entire specimen (in the field of view of the objective), it excites all the fluorescent molecules in the entire volume of the specimen. Thus, light emitted from the entire volume will be detected. This may create a background fluorescence that will affect the overall signal to noise ratio.
Several solutions have been developed, among them Confocal microscopy, TIRF and others which we will discuss in later posts.
Green fluorescent protein (GFP in short) was the first of a large (and still growing) family of proteins with the unique ability to fluoresce in different colors. But since GFP was the first, and it is still the most popularly used, and most known by the general public, I dedicate the first post to this protein.
But first, let’s briefly define the term fluorescence: it is the emission of light by a substance that absorbed light. The emitted light is at a longer wave-length than the exciting wave-length. However, under certain conditions where the fluorofore is simultaneously excited by two photos, the emission is shorter than the exciting wave-length. We will get to two- (and three-) photon excitation at a later post. Fluorescent is different than phosphorescent – which is emission of light independently of any excitation.
The common laboratory GFP is excited by blue light and emits green light.
But what is GFP?
GFP is a natural protein that was first isolated from the jellyfish Aequorea Victoria in 1962. Although 50 years has passed since its discovery, the biological function of GFP and GFP-like proteins remains controversial. I may dedicate a post about it at a later time.
GFP is a 238 amino-acid long protein, with a unique barrel-like structure. Unlike many proteins that utilize co-enzyme molecules to elicit their function (e.g hemoglobin that utilizes the heme molecule), the uniqueness of GFP is of creating its own chromofore by cyclization of amino acids number 65-67 (serine-tyrosine-glycine). This self-assembly is one of GFPs advantaged that made it so popular.
Another important feature of GFP is its monomeric, i.e. each protein functions alone, and does not associate with other GFP proteins (except when the concentration is high). There are other fluorescent proteins that act as dimers (i.e. two proteins) or even tetramers (4 proteins).
The wild-type GFP from the jellyfish is accustomed to low ambient temperatures (since the jellyfish is found in the cold Pacific Northwest). Therefore it has a low efficiency of folding at 37°C, which is required for studying many biological systems, from bacteria to mammalian. Another deficiency of the wild-type GFP is its low fluorescence intensity after excitation with blue light. To improve the quality of the protein for research, two mutations were implemented. The first is S65T (serine 65 changed to threonine). This made the protein fluoresce 35 times brighter than the wild-type GFP. The S65T version is often used in systems at low temperatures (20-30°C) such as insect of yeast. The second mutation, phenylalanine 64 changed to leucine (F64L), improve the folding of the protein at 37°C. GFP protein with the both mutations is called enhanced-GFP (EGFP in short). In the following posts, I may use the term GFP, instead of EGFP or GFP(S65T). In any case, few people still work with the wild-type GFP protein.
The above mutations also changed the spectral properties of the GFP protein. The wild-type GFP has two excitation peaks: a major one at 395nm and a minor at 475nm. If the GFP is excited at 395nm (UV light), it emits green light at a wavelength maximum of 508nm. Excitation at 475nm gives a maximum of 503nm. The S65T mutation leads to chromophore ionization. The excitation at 395nm is suppressed (due to the neutral phenol of the threonine) and the 475nm excitation peak is shifted to 488-490nm and enhanced 5-6 times.
Thus, you will find in most fluorescent microscopes a light source at 488nm, to suit the excitation peak of the commonly used EGFP.
There are several other classes of mutations that were introduced in GFP in order to create other spectral properties (e.g. red or blue-shifted, photoactivatable GFP).
So, how is GFP used as a fluorescent marker in biological research? Simply put, you can very easily fuse GFP to any protein or peptide of your choice. In some cases, this may disrupt the function of that protein. But in many cases the disruption is minimal if any. Once the GFP-fusion protein is expressed in the cell, you can answer, by using fluorescent microscopy, the following questions: WHERE in the cell does it reside? Does it MOVE? IF and/or WHEN is it expressed and what is the LEVEL of expression? What is its rate of SYNTHESIS or DEGRADATION? WHO does it associate with? And many more questions which we will explore in this blog.