Summary Microscopy (CLB-30306)
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Contents
Basics of microscopy: .................................................................................................................................................. 2
The NA (Numerical Aperture) .................................................................................................................................. 2
Lenses ..................................................................................................................................................................... 2
Diffraction and resolution ....................................................................................................................................... 3
Image contrast generation .......................................................................................................................................... 5
Brightfield microscopy ............................................................................................................................................ 5
Dark field microscopy.............................................................................................................................................. 5
Phase contrast microscopy ...................................................................................................................................... 5
Polarized light microscopy....................................................................................................................................... 6
Differential Interference Contrast (DIC) microscopy ................................................................................................ 6
Types of microscopy: .................................................................................................................................................. 7
Compound Microscope: .......................................................................................................................................... 7
Fluorescence Microscope: ....................................................................................................................................... 9
Optical Sectioning Techniques: .............................................................................................................................. 10
Super-resolution Microscopy .................................................................................................................................... 12
Single Molecule Imaging ....................................................................................................................................... 12
Photoactivated Localization Microscopy (PALM) ................................................................................................... 12
Stimulated Emission Depletion (STED) ................................................................................................................... 12
Reporter proteins ..................................................................................................................................................... 13
Inserting reporter genes........................................................................................................................................ 13
Fluorescent sensors .............................................................................................................................................. 14
Fluorescent Labels ................................................................................................................................................ 16
Light Activated Experiments ...................................................................................................................................... 17
FRAP ..................................................................................................................................................................... 17
Photo activation .................................................................................................................................................... 17
Uncaging ............................................................................................................................................................... 17
Fluorescence Lifetime Imaging (FLIM) ................................................................................................................... 17
Basics of microscopy:
The lens equation: 1/f = 1/s + 1/s’
Magnification: y’/s = s’/s
• F: focal length
• s and s’: object and image distances
• y: object size, y’: image size
To calculate your magnification:
..x objective lens * ..x eyepiece lens = magnification
..mm focal length tube lens / ..mm focal length objective lens = magnification
The NA (Numerical Aperture)
The NA is a measure of the angles of light that can be
collected by a lens (max of 1 when no immersion
medium is used). Higher NA lenses collect more light and
form less noisy images.
NA = n sin μ where n is the refractive index of the
medium and μ is half the angle of the collection cone.
Lenses
Immersion lens: water or oil immersion increases the numerical aperture
Rays coming from the object may become reflected between the coverslip
and air. These rays are not collected for image formation, but liquid
immersion prevents this.
Oil: n=1.516. So, a lens with NA = 0.95 will result in a 0.95*1.516 NA = 1.45
Plan designation lens: corrected for spherical aberration
Spherical aberration is when parallel light rays that travel close and far away
from the optical axis are not focuses to the same point. Therefore, the
image appears blurred.
Achromat objective lens: correct for chromatic aberrations
A lens with chromatic aberration is made of a material with a wavelengthdependent index of refraction. Therefore, it refracts different colours
differently
So, plan achromats are corrected for both spherical and chromatic
aberrations.
Image brightness increases with the square of the NA value of the objective
lens, increases with the use of oil-immersion, and decreases with image
magnification.
Diffraction and resolution
Diffraction is the bending of light waves around small objects and openings like water
waves. For this fundamental reason there is a limit to the size of individual features that
can be distinguished. A parallel beam forms a diffraction-limited airy pattern, which is
about 200 nm in size. The radius of the first dark ring in the airy pattern, and thus the
resolution, equals RAiry = 1.22 * λ / (NAcondenser + NAobjective). For top quality lenses this
equals about 1.22 * 530 nm / (1.45+1.45) = 223 nm. The better the NA, the better the
resolution. The resolution is inversely proportional to the wavelength, so small
wavelengths like blue light give a good resolution.
The Rayleigh criterion:
Two objects can just be resolved if
the spacing is equal to the radius of
the airy disk, which is about 200 nm
Pixel size
The optimal spacing of pixels is; not too large to cause loss of detail, but not too small to collect inadequate signal.
The Nyquist criterium states that the pixel size that optimizes signal per pixel, whilst maintaining fine resolvable
detail is around half the radius of the airy pattern (half the resolution of a microscope).
If your pixels are too small, a process called pixel binning can be used to sum the signal of a group of pixels. It will
however decrease resolvable detail.
Koehler illumination:
Kohler illumination illuminates the sample evenly and allows control over illumination angles. The light source is
focused by an additional lens in the focal plane of the condenser lens. This way each point of the lamp generates a
parallel beam of light that illuminates the whole sample but under a different angle. Closing the condenser aperture
decreases the range of angles at which the sample is illuminated.
Field diaphragm and condenser lens
The field aperture controls the size of the illuminated area
The condenser aperture selects the angles under which the sample is illuminated
Image contrast generation
Brightfield microscopy
This type of microscopy relies on the local change of
colour/intensity of pigmented areas in the final image. In
general, the contrast is low, but this can be improved by using
histological stains.
Dark field microscopy
In dark field microscopy the sample is illuminated with a hollow
cone of light because a stop in the back focal plane of the
condenser blocks light under small angles. If no sample is
present the rays will continue in a straight line and no
illumination will reach the objective lens. Only light that is
scattered by a sample will contribute to image formation.
A disadvantage is that this technique is very sensitive to stray
dust and other artefacts that deflect light.
Phase contrast microscopy
Illumination with a hollow cone of light will be scattered by a
sample which causes about ¼  phase shift change. A phase
plate in the objective lens imposes an additional ¼  phase shift
on the scattered light only. Destructive interference then
locally darkens the image to generate contrast.
This technique is often used for imaging tissue culture cells as it
visualizes “phase dense” objects. Limitations are a halo around
the specimen, and its not usable for thick specimens.
Polarized light microscopy
A first polarizer selectively passes only one angle of polarization. A
second polarizer rotated 90 degrees will block this light completely.
Birefringent (the angle of light refraction depends on the polarisation
direction of the light) samples however can rotate the polarization
axis, thus an image is formed.
Differential Interference Contrast (DIC) microscopy
DIC combines concepts from polarization and phase contrast microscopy. Samples do
not need to be birefringent. The technique detects a phase change that is generated
between two light rays that take a slightly different path through a sample. A phase
change is generated when one of the rays travels through an object with a higher index
of refraction. This phase change is converted to a change in direction of polarization by
a Wollaston prism. This change in polarization is then detected similarly as in a
polarized light microscope.
Because the phase delay can be by either of the rays
the phase difference can result in both a higher and
lower intensity. So as a result the image has a shadow
cast appearance with dark and bright edges
specifically at boundaries of different materials.
Summary:
Bright Field Microscopy:
Simplest form of microscopy, is frequently used in combination with histological stains to generate contrast.
Dark Field Microscopy:
It uses the phase of light to generate image contrast. It requires the NA of the objective lens to be smaller than the
NA of the condenser lens.
Phase Contrast Microscopy:
It relies on destructive interference of scattered and unscattered light rays with different phases. It relies on a phase
shift of 90 degrees when light Is scattered.
Polarized Light Microscopy:
Generates contrast on samples that are birefringent.
DIC Microscopy:
Generates contrast depending on the size and direction of the local gradient in refractive index within the sample. It
uses two Wollaston prisms to generate and re-combine two spatially-shifted light rays that are orthogonally
polarized. Wollaston prisms are only used for DIC microscopy.
FRAP Studies: the rate of Fluorescence Recovery After Photobleaching.
Types of microscopy:
Compound Microscope:
It uses a two-step magnification where the total magnification is Mobjective x Meyepiece .
The field aperture’s function is to prevent stray light (photobleaching) by limiting the illuminated area of the sample
to the size of the observed area (field of view).
A condenser lens collects and focusses light from the illumination source onto the specimen.
Infinity optical system:
It uses two lenses but there is no intermediate image. The object is placed in the focal plane of the objective. This
way each point of the object forms a parallel bundle of light. A tube lens forms a real image on a camera sensor
where the magnification is ftube / fobjective . In the infinity space of parallel light bundles auxiliary optical components
can be introduced without much effect on the image formation.
Fluorescence Microscope:
A molecule is placed in an excited state by absorption of a photon. When the
molecule decays back to ground state a photon with less energy is emitted (less
energy = longer wavelength = red shifted). This shift in wavelength equals the
Stokes’ shift.
A basic fluorescence microscope needs a light source that emits colours, an
excitation filter that selects a usable wavelength, and an emission filter that blocks
the excitation light and passes fluorescence light to the camera.
Fluorescence’s advantages are; sensitive, specific, multiparameter (intensity,
bursts, polarization, lifetime), living cell and dynamics. Disadvantages are; labelling
and introduction into the cell, phototoxic effect, sample thickness and fluorophore disturbance.
Different light sources:
Halogen/Tungsten lamp: light intensity is distributed over a continuous spectrum. Excitation filters are used to select
specific excitation colours.
Gas discharge lamp (Mercury, Xenon arc): light spectrum contains intense peaks at certain wavelengths.
LEDs: Narrow wavelength bands can be produced, but each fluorophore requires a different specific LED.
LASERS: High intensity in a single wavelength, so no filter required to block unwanted colours.
Epi-fluorescence microscope:
Excitation light does not shine in your detector and background light is
efficiently blocked. The objective is now used for both illumination and
observation
Resolution
= Dmin = 1.22 * λ / (NAcondenser + NAobjective)
= Dmin = 1.22 * λ / 2 NAobjective
The limitation of fluorescence is the inevitable photo bleaching that
takes place due to excitation. Besides bleaching, also quenching
decreases intensity of fluorescence. Quenching is the transfer of energy
to nearby acceptor molecules through various mechanisms.
The thickness of an in-focus slice is equal to the z resolution of a
microscope = dz = 2λn/NA2
Optical Sectioning Techniques:
Confocal Laser Scanning Microscopy (CLSM)
The principle was proposed by Minski (1955) and encompasses the use of spatial filtering techniques to eliminate
out-of-focus light. By placing an aperture (pinhole) in the image plane, the out-of-focus light cannot reach the
detector. This enhances the signal to noise ratio (enhanced contrast), allows for better sectioning (thinner slices in zdirection) and improves axial and lateral resolution.
In a conventional wide-field fluorescence microscope a mercury lamp illuminates the entire specimen. A confocal
microscope uses a parallel laser beam that is focused into a small diffraction spot. Scan mirrors in the excitation path
can move this spot for point-by-point measurements. In this way an image is generated of the fluorescence emitted
from a horizontal plan at a certain depth inside the sample, called optical sectioning. Confocal microscopy can create
3D-images of the structure but also show dynamics of cellular processes.
Image Deconvolution
A stack of images is taken at different heights, which are combined whilst applying
deconvolution to redistribute image intensity over the image stack to remove out
of focus blur.
The drawback is that fluorophores bleach quickly. Even if only a single plane is
required one still needs to acquire a z-stack to apply the deconvolution algorithm.
Also does not work with life cells because of movement during imaging.
Spinning Disk Microscopy (SDM)
A disk with microlenses focuses an excitation beam onto the holes in
the second disk; the pinhole or Nipkow disk. An array of pinholes on
the Nipkow disk are spaced apart just enough such that fluorescent
light generated does not mix with neighbouring holes.
Each location on the sample is illuminated about 1000 times per
second, so illumination takes place not once but multiple times
during a camera exposure of roughly 100ms.
Compared to CLSM there is a lower local excitation intensity, so
bleaching is minimized. It is however less flexible for FRAP/FRET and
has a fixed pinhole size.
Multi-photon Microscopy
Instead of using one light source with a certain wavelength to excite a
fluorophore one can use two lower-energy photons at half the energy
(e.g. 2x 1000nm vs 1x 500nm), but only if the two photons arrive
nearly at the same time. Such an event’s likeliness increases with the
square of the excitation intensity.
The advantage is that higher
wavelength penetrates deeper
into tissues compared to
visible light. No excitation and
photobleaching takes place in
out-of-focus planes. You do
however need more complex
pulsed lasers that deliver high instantaneous intensity. Also, photobleaching is
accelerated in the in-focus plane.
Total Internal Reflection Microscopy (TIRF)
A sample is illuminated through the objective by a shallow
angle light ray that reflects at the cover glass – water interface.
Above a critical incident angle (about 62 degrees) light is fully
reflected. An evanescent light field however penetrates about
100 nm into the water. Consequently, only fluorophores near
the interface are excited, and high contrast is achieved because
there is no fluorescence from other planes in the sample.
Selective Plane Illumination Microscopy (SPIM)
One plane of the sample is illuminated by a thin light sheet
coming from the side. Little out-of-focus light is present
because other planes are not illuminated. The technique can be
combined with image deconvolution to further increase
resolution.
Fluorophores in other planes do not photobleach.
Summary:
Super-resolution Microscopy
Single Molecule Imaging
Whilst it is not possible to resolve single molecules that are close
together, it is possible to localize single fluorophores precisely if their
spacing is sparse.
Emitted light is distributed over several neighbouring pixels, but the
centre can be estimated with an accuracy smaller than the individual
pixel size. Up to 1nm accuracy can be achieved if sufficient photons are
detected. The theoretical position accuracy =
Diffraction limited resolution / √N
Photoactivated Localization Microscopy (PALM)
Fluorophores like the photoactivatable PA-GFP can be switched on by a
specific wavelength. Single activated fluorophores in the on-state are
detected with light of a different wavelength, fluorophores in the offstate are not visible. The temporarily activated fluorophores are imaged
and their positions can be determined with ~20nm accuracy. The overall
image is reconstructed with a computer from the positions obtained
from hundreds of imaging cycles. This however requires the sample to
be completely immobile, but faster cameras improve on this.
Stimulated Emission Depletion (STED)
Fluorophores can be stimulated to decay quickly to their
ground state using light at the emission wavelength. In STED
microscopy fluorophores are first excited in a confocal laser
spot. Directly after this the confocal volume is illuminated with
a doughnut-shaped beam profile after which only the
fluorophores in the centre of the confocal volume remain in
their excited state. These fluorophores decay and emit light
which is detected. Because the detected light originates from a
spot smaller than the diffraction limit, up to a 10-fold
improvement in resolution is achieved. The setup is
complicated however, and photo bleaching is rapid.
Reporter proteins
Colour stain as reporter
-> enzyme
E.G. The bacterial lacZ gen with blue pigment
Bioluminescence reporter
-> enzyme
E.G. The bacterial luciferase (LUX), the firefly luciferase (ffLUC) or Aequorin
LUC is good for promotor dynamics
Fluorescence reporter
-> structural protein
E.G. Green Fluorescent Protein (GFP) or tdTomato
GFP is good for cellular resolution promotor activity, subcellular localisation studies or protein stability
studies.
Inserting reporter genes
Transient transformation (100-1000000 gene copies)
This quick but temporary implementation is transitory and will thus only last a short time.
Agroinfiltration;
The leaf is infiltrated with activated A. tumefaciens solution carrying the reporter construct. The pathogen can
transfer Ti-DNA to cells of the infiltrated tissue. The Ti-Dna is expressed in the cells but will be degraded after a few
hours/days.
Particle bombardment;
Gold particles are coated with DNA containing the reporter construct. The plant tissue is bombarded with the coated
particles. Cells that survive can express the introduced DNA
Transfecting protoplasts;
Protoplasts are plant cells without a cell wall. The protoplasts are suspended in a solution with the reporter DNA
construct. Temporary disruption of the plasma membrane via chemical treatment, electro or heat shock will allow
many copies of the reporter DNA to diffuse into the nucleus of the cell and be transcribed.
Stable transformation (1-10 gene copies)
Integrating the reporter gene into the genome is time-consuming but it has advantages. It has both the ability to
retain a reporter gene for multiple generations, and it also has the ability to cross in order to generate double
labelled plant lines.
Agrobacterium tumefaciens is a plant pathogen
that injects its Ti-plasmid into the plant cell. It
integrates at random into chromosomal DNA to
reprogram plant cells. By engineering the
pathogen, it is possible to insert modified pieces of
DNA. Insertion is random however, so your
construct can disrupt genes if integrated at the
wrong position.
Heterochromatin is tightly packed DNA, Euchromatin is loosely
packed DNA. Integration into heterochromatin results in no
transcription. This can however be prevented by using Matrix
Attachment Regions to flank your reporter gene which
connects chromatin loops with histones.
Fluorescent sensors
FRET based biosensors:
FRET: Föster Resonance Energy
Transfer is the transfer of energy
from an excited chromophore to
a neighbouring chromophore.
FRET is good for interaction
studies between two binding
domains that alter spacing by a
binding ligand, or for two
proteins that interact. The
energy transfer is non-radiative
and through space.
The prerequisite for FRET is R0 = the critical distance between donor and
acceptor where energy transfer is 50%.
R0 = 0.0211 x (K2QD J n-4 )1/6 (nm)
K2 = the dipole orientation, when random it’s 2/3
QD = the Quantum yield of the Donor (0<QD<1)
J = the spectral overlap between donor emission and acceptor
absorbance
N = refractive index (1.4 inside fixed cells)
The efficiency transfer is inversely proportional to the
sixth power over the distance R between the Donor and
the Acceptor.
In practice the formula to calculate FRET =
If-BTd*Id-BTa*Ia
If = Intensity FRET channel (acceptor channel) with donor excitation wavelength
Id = Intensity Donor channel with donor excitation wavelength
Ia = Intensity Acceptor channel with acceptor excitation wavelength
BTd = Bleed through from donor
BTa = Bleed through due to direct excitation of acceptor
FRET by acceptor photo-bleaching
Calcium sensor
Aequorin + Calcium generates an excited aequorin protein. By coupling a GFP to
aequorin energy is transferred from aequorin to the GFP, emitting green light. As
such aequorin-GFP can act as a calcium sensor.
Bimolecular Fluorescence Complementation (BiFC)
The association of fluorescent protein fragments that are attached to components of
the same macromolecular complex.
Auxin Sensor DR5
Auxin is a plant hormone with response factor ARF.
ARFs function by binding auxin response elements,
thereby changing transcription. A site directed
mutation of AuxRE = DR5. This DR5 can be coupled
to form DR5:GUS / DR5:GFP auxin reporters
Green Fluorescent Protein
Can reveal both promotor activity and protein localization.
Potential GFP artefacts:
If GFP protein blocks the signal peptide there is no normal localisation.
Expression level of protein should not saturate transport routes within the cell. GFP is about 240 amino acids in size
so it might change diffusion/transport of the target protein. It might also block the catalytic centre of the target
protein.
Fluorescent Labels
Intrinsic fluorophore
+ no labelling & purification steps
+ no extension of molecule
- low quantum yield
- sensitive to photobleaching
- selectivity
Chemical dye
+ high quantum yield
- labelling & purification steps
- sensitive to environment
- sticking to intracellular components
Green fluorescent protein
+ ‘genetic’ labelling, no purification steps
+ high quantum yield
+ protected from environment
- size
Quantum dots
+ photostable
+ different beads excited at one wavelength
+ sharp emission bands
+ high extinction coefficient
+ size
Ratio-probes
Easily implemented
Fast (suitable for live-cell imaging)
Accurate spatio-temporal information
Not quantitative (relative probe concentration)
Independent of:
Probe concentration
Light intensity
Photobleaching
Light Activated Experiments
FRAP
Fluorescence Recovery After Photobleaching is a technique to look at
fluorescence recovery after photobleaching in a subpopulation of molecules.
Fluorophores within a small region are quickly photobleached with a focused
laser beam. The subsequent exchange of fluorophores with the surrounding
non-bleached region is imaged using low laser power.
Apart from diffusing, molecules may reversibly bind to cellular targets. E.G. a
fluorescently-tagged transmembrane protein at the nuclear envelope binds to
the lamin cytoskeleton that supports the membrane. The mobility of the
transmembrane protein is now jointly determined by the average lifetime of
the bond to lamin and its diffusion speed in the membrane.
If the timescale of diffusion is fast compared to the timescale of protein-target
dissociation, then the fluorescence recovery timescale is a direct measure for
bond (off-rate=1 / recovery timescale)
Photo activation
A mutated version of GFP can be photo-activated by UV light. The excitation spectrum changes after activation and
green light is now emitted upon blue light excitation.
Uncaging
With uncaging a covalent bond is broken by UV light. This results in an
activated form of the molecule. After uncaging the cellular response to the
activated molecule can be imaged. An example is ATP.
Fluorescence Lifetime Imaging (FLIM)
Relaxation of excited molecules take place with a certain probability based on the radiative and
non-radiative decay rates. Different molecules have different exponential decay rate of
fluorescence, which is the basis for FLIM. Fluorescence lifetime imaging is based on differences in
excited state decay rates from a fluorescent sample. The advantages are that the measure is absolute, an intrinsic
property, independent of probe concentration or laser intensity, and it’s sensitive to the environment.