Advanced microscopic techniques and nanotechnology, inglese, Microscopy, Medical biotechnology
Typically in another box there are one or more lasers and you can shine one or
more colours at the same time because sometimes is useful to look at more
than one fluorophore at time; so you need to be able of scanning but you also
need to be able to separate that by colour so what do you need to do? Instead
of just a photomultiplier (PMT, detector that just measure the light that arrive,
no matter what colour is it), I put something more to separate the colours.
To separate the colours, I just use another box that is able to read the intensity
of light as function of the colour because it has inside a diffraction grate. So
light arrives and it’s separated in colours and it’s sent all together to the
detector, a multi anode which is like a system of different eyes/detector and
each detector gets a range of each colour. enough light
You can do this if you have
because you’re illuminating a point in the
The question is: when the light goes back how
much light goes back?
Usually you use a PMT because you receive
very little light so you have to be very
After computing you have a 3D image
because it has
Multibeam + CCD
Instead of scanning with just one beam, I can scan with many beams together,
if I have a source of light which is powerful enough.
If the light is powerful I can decide to separate it in more points at the same
time. How do I do that? I can use a Nipkow disc, it’s a wheel with many holes
If the wheel rotate you can see through the holes, and in a complete turn you
had all the holes per line. 9
Then if you have a lamp on the other side and if you can synchronize the things
you could have that your hole is moving and there’s a light here but the light
that gets intense or dim as the hole moves so on your retina you get a line
which is bright-bright-dark-bright dark etc.
If the wheel turn fast enough the light remain on your retina and you have an
image (is like the old TVs before the catotube).
You have a light coming from a laser (you
don’t focus it), you send on the first disk
and this disk has a lot of small lenses
that focus the light on a second disk of
pinholes: the two disks are attached to
each others so they move together.
So when the light encounters the first
disc is divided into different beams, each
lens focuses on a different place on the
disc with the pinholes and goes through
Passed the pinholes, you’re exactly in
the situation that you had before
(objective and the sample) but it’s like if
you have different lasers.
The light that goes back from the sample goes through the pinholes as before,
what pass through the pinholes is the light on the focal point of the beam, not
much light from below or from above. Then with a partial reflecting mirror you
can send it on a very sensitive camera (CCD).
By rotating the disc you can move the spots on the sample, so you can see
lines, and as you turn you fill the space, until in one turn you have filled the all
With multi beam confocal microscopy you can go really fast, it is very useful
when you are in a hurry and you have to see something that is happening and
you have to take an image before it goes.
There’s a lot of problem also in communicating data because you can go really
fast but you also have to transfer this data to the computer, the speed of
communication is crucial.
There’re some problems:
- You can go fast but you don’t have enough light
- You can go fast and take a lot of images but it’s not clear where you put
them, so if you want to go really fast you have to use local memories
- Incredible amount of data produced
Intrinsic concept of scanning microscopy
Pixel is a picture element, is the size of unit surface by which I divide an
Pixel is an intrinsic unit because when I’m scanning I divide the space in
positions, I don’t have a continuous result. The system says I’m in position 1,
there’s intensity of 10, then the system move to position 2: I’m dividing space
in distinct positions.
So to get an image you have to say position and light intensity in that position.
If it’s in 2D is like a matrix, I’m dividing in squares and each square is called
pixel. xy, z
But here you don’t have only I also have so I have to divide the position in
xy and scan all the positions. So my unit element is not like in the picture a
square, it’s a volume because has and is called Voxel.
This makes impossible to see the image without a computer, because you have
a list of position and a list of intensity.
One thing is how do I divide the space into positions (Voxel) and another thing
is the minimum volume I can see as a resolution: it’s called Resel and, I cannot
distinguish anything inside, because this is the minimum thing that I can see.
Voxel and Resel are not the same thing, one is intrinsic of the optics which is
the minimum volume element I can resolve and the other is how do I decide to
split the space (it can be 100 times larger or 100 times smaller than
resolution), I decide it telling the piezo system where to move the beam.
I want to have Resel and Voxel to coincide so that each position is a resolution
element, but the fact that they coincide is not intrinsic.
Of this cylinder I know how high and how wide it is, it’s not that within I cannot
seen anything and without it’s clearly distinguished. When I shine laser into a
point I can distinguish that my light is coming mainly from the cylinder but is
also coming a little bit from the side too (is a continuous function).
In confocal microscopy I have and overlap of Resels from voxels, each voxel
should contain a Resel but contains also light that is coming from the neighbour
Resels; so the light getting from a position is contaminated a little from the
light of Resels around, because the resolution is not sharp (even if I make
Resels and Voxels coincide).
So what I can do? I can correct a little bit but if I know the shape of my Resel
(which means if I know the shape of my point spread function) I can take this
image, do the Fourier’s transform and then there’s a filter that I can use in the
Fourier’s transform to correct it and then go back to the image.
The problem is that sometimes, pixel or voxel of an image are contaminated by
neighbours and there is trick called that can partly clean this
Confocal microscopy advantages
- You get at the limit of resolution, it’s easy to clean the system but you
don’t go very further other microscopy
You can have 3D images
- (most important aspect) what you get is a
composition of the image.
Doing scanning, if you have light enough you can go faster or you can look at
colours: the output are images where you can get different coloured images;
for example you can get an image where you can split the image in a portion
with a wavelength and a portion with another wavelength.
If you have a system with 2 fluorophores if you put together you’ll see just
black and white, you don’t distinguish colours, or you can split in 2 regions and
you have 2 images and you can give false colours to the fluorescent signals. If
you have fluorophores that are well separated you have a good situation, but
sometimes you have fluorophores that have an emission spectra overlapped
and in this case you need more information, is hard to have a situation in which
you can split the colours.
You can have a system of images and which of this is distinct by different
wavelength: we can have a stack of images, each of these is a different
position of depth for example, but we can also have a stack of images each of
these is a different wavelength, so if you want to put them together you’ll have
a 4 dimension object and .
If you have a signal grey made by red and green fluorophores, if you know the
spectra of the 2, there’s a mathematical procedure by which you can know how
much green and how much red you have. What I get is a mix of the 2 so I need
to unmix them, but to be able to unmix them you need to know the behaviour,
the value as a function of different values, of fluorophores.
I need to separate the colours either because we have very well split
fluorophores and it’s easy or because you have a mix of fluorophores, in that
case you have a stack and for each position you can tell how much you have
green and how much you have red: so for each voxels you can separate the 2
Light Sheet Microscopy
It’s a kind of variant of confocal microscopy.
I can look at the whole picture, as I did before, but now I can illuminate only
a sheet/plane in the direction: so it’s a kind of combination because I’m
looking in 2D but the system is illuminated only on a layer, so I’m getting the z
resolution because I’m illuminating only a plane.
You can illuminate the whole plane or you can use a beam that scans through
the all plane. You illuminate from the side and look through the objective.
Selective Plane Illumination Microscopy (SPIM)
It’s called or Light sheet.
It remains with the resolution limit of all the other microscopy.
L i g h t s h e e t m i c r o s c opy
E In light-sheet microscopy, fluorescence excitation (blue arrow) and detection (green
In light-sheet microscopy, fluorescence excitation (blue arrow) and detection
R arrow) are split into two distinct optical paths.
The illumination axis is orthogonal to the detection axis and is designed to illuminate
(green arrow) are split into two distinct optical paths.
a very thin volume around the focal plane of the detection objective.
Many different implementations of this principle exist, however, the most common
The illumination axis is orthogonal to the detection axis and is designed to
one is the generation of a sheet of laser light that illuminates the sample in the focal
plane from one side.
illuminate a very thin volume around the focal plane of the detection objective.
Many different implementations of this principle exist, however, the most
common one is the generation of a sheet of laser light that illuminates the
sample in the focal plane from one side.
While in confocal you illuminate point by point, through the pinholes, and so
you’re sensitive only on a plane; in light sheet you’re looking all together but
you’re illuminating just one sheet, so the detection is all together on one plane.
- Not complicated as the confocal
- Less photobleaching (you broke the fluorophores only where you’re looking,
in confocal you shine also above and below but you’re not sensitive to these
point). 5 Ottobre 2016
What do you want to avoid with bleaching? You want to illuminate only what
you want to look at. When you use a confocal microscopy you illuminate also
the below and the above portion of what you are focusing on. With the light
sheet microscopy you only illuminate what you look, just the plane of interest
(but you look at the hole system of fluorophore).
Is it possible to overcome the resolution limit set by diffraction?
Both 2D microscopy and scanning microscopy (as confocal) obey the diffraction
resolution limit. image
The resolution applies as we make a linear image of an object. By we
mean that we use a lens to make an image; linear means that the optical
signal you’re looking at, is proportional to the intensity of the light that is
light transmitted or emitted by the molecules in
shining, so it means that the
the sample is proportional to the light that is used to illuminate or excite them.
To overcome the diffraction barrier you have to overcome one of these two
concepts, or both:
By inducing the sample by a proper choice of molecules illumination,
non-linear mechanisms of absorption or emission
By replacing the use of optical images with other strategies.
There are several types of scanning microscopy:
- scanning with production of an image
- SEM (Scanning Electron Microscope) scanning with production of an
AFM (Atomic Force Microscopy)
- scanning without making an image
If we want to make something in the middle between confocal and atomic
force, we could think about a stethoscope that makes a sort of a bigger image
of the heartbeat. 13
We could make a stethoscope with light: the NSOM/SNOM (Near field
Scanning Optical Microscopy) is a type of optical scanning
generation of an image.
It’s a capillary with an optical fibre in the centre (70 nm): you can go and put
your fibre tip in the sample that you want to look without making an image,
it doesn’t use lenses but it just collects light from that point of interest.
So the tip goes near to where the light comes out and collects the light from
I have this tip and I can:
- Illuminate the sample and look for the light emitted by the fluorophore in
- Shine and collect light from the same source locally.
I just have to be able to mechanically bring around the tip: for example I can
touch the membrane of a cell with the tip!
I can go beyond the diffraction limit
So with NSOM/SNOM , because if I don’t
make an image, I can go smaller than the wavelength!!
Two photons excitation microscopy
It’s a kind of not linear microscopy in which you can excite fluorophores with
two photons. In the graph, there are the energetic levels of
molecules (black straights): the molecules have
different levels because of vibration; the lower
level is the electronic ground state.
In violet there is the energy of a molecule which
is excited one time.
During fluorescence you shine light on a molecule
and if the molecule carries enough energy then
the system absorbs this energy (360 nm) and
goes to a higher vibrational state.
But when the vibrations are quenched, the system goes back to a lower state.
If the molecule absorbs 360nm, when it emits light, it will emit at a lower
energy (460nm) because it has lost part of the energy because of vibration.
So in fluorescence molecules absorb a certain quantity of energy and release a
smaller quantity of energy: since the quantity of released energy is smaller
wavelength is longer (460 nm instead of 360 nm).
In the two photons excitation microscopy you can excite the same fluorophores
by using simultaneously two photons that have half of the energy (720nm): the
at the same
photons have to arrive time on the same molecules, or within a
very short time.
We can do the same thing with three photons too.
Normally the absorption is proportional to the intensity (I ). If the photons
are two, the amount of energy absorbed is no longer proportional to the
intensity but to the intensity squared (I ), because you need two photons
to be at the same time and hit the same molecule. This means that you need
more photons because they have to be in the
same place at the same moment.
In linear absorption the intensity of excited
fluorophores is proportional to the intensity of
shining light while their number is proportional
to the intensity of energy: the total energy is
but the intensity varies
always the same,
through the hourglass.
number of fluorophores excited is always the same the density is
So the but
greater in the middle. linear absorption.
This is what happens with
The profile of the intensity of the beam is like a
Gaussian and this is also the profile of the intensity of
the excited fluorophores, if the probability is
proportional to the intensity (as a single photon).
In two photons, if the probability is proportional to
the squared of the intensity, the probability of
much larger in the centre
excitation changes: it is
and less in the side, much more
so it becomes
probable to excite the fluorophores in the centre of
With double photons emission I’m concentrating the excitation in the centre of
the beam,: with this technique only molecules in the centre of the beam get
excited and so I narrowed the region where I can excite molecules!
lose resolution because I use a bigger wavelength,
However using this method I
using a smaller portion of a larger beam.
What do we gain? When I shine light with linear excitation, I’m exciting photons
everywhere where the beam goes, while if I have two photons I’m just exciting
where the beam is minimum without the
need of having a pinhole (I’m doing a sort of
The other thing I gain is that I have much
less photobleaching because I’m exciting
only the centre of the beam so I’m not
wasting fluorophores below and upper, so
the fluorescence I produce is just where I’m
looking at. Moreover I use longer
wavelengths so I can penetrate better the
much powerful sources,
For this method we also need to use like a million
times more powerful. It is also possible to use pulsed sources that don’t shine
photons continuously but accumulate photons and the shine them all together
in order to have a very concentrate energy within the pulses.
STED (Stimulated Emission Depletion fluorescence)
In order to have absorption I have to shine light, while to have an emission I
don’t have to shine light.
molecule to emit energy if I shine it whit the
I can convince a
same light that it emits: for example if I shine my molecule in
green but also in red (that is my emission light, as in figure),
the energy decay become faster. Not only I release my light, I
release it in the same direction and with the same maximum
stimulated emission is the phenomenon by
which the rate of emission of an excited molecule increases
proportionally to the intensity of light that it is illuminated by, if
the light is the same wavelength of the fluorescence emission
STED uses two lights:
- One for the excites the fluorophores
- One for the makes the fluorophores de-excited.
The number of molecules which
are on level 2 (dn ), the energy
from which we started to get
fluorescence, change in time by
virtue some terms one of that is
proportional to the intensity of
light that shines (h ) times the
difference between level n and
level n .
If level n is more populated than level n light comes and goes down.
If level n is more populated than level n the light can also go back.
(formula riquadro rosso).
I can de-excite the fluorophores with light.
So in STED I’m using two beams of two different shapes:
excitation pulse normal
- The is a focus beam
de-excitation pulse donut-shape
- The is a beam darker in the centre and
bright on the side.
So I excite molecules were the beam is, then I turn down the fluorophores
around with the de-excitation beam. 16
Non linearity is used in de-excitation and
to do that you have to shine stronger and
Increasing the power of light, the hole
become smaller and smaller because the
probability to getting de-excitated is not
Since the stimulated emission is brought
in a non-linear regime when you increase
a lot the power, the region where the
fluorophores are not de-excited becomes
really small: increasing intensity the light areas become saturated and dark
areas are contracted. We gain much if the turning off is proportional to
the intensity: I have to use non linearity in de-
excitation, so in turning fluorophores off and I can
do this shining stronger, with super powerful pulse.
With this method of increasing power the hole
becomes smaller and smaller: the probability of
being de-excited is proportional to the de-
excitation pulse when this pulse is not very
powerful; because the probability of being de-
excited cannot be more than a limit, when the de-
exciting pulse goes over this limit, all the
molecules become de-excited.
*Dashed lines indicate the percentage of quenched
fluorophores.Upon entering the non- linear regime
the region where we can still have excited
The much I increase the power of the de-excitation light, the narrower becomes
the region where I still have fluorophores emitting light
*the dashed blue line represents the 100% quenching level.
This technology is made up of a first green excitation laser and
then a second read beam that has a donut-shape; the size of the
hole is the resolution; the red pulse is much stronger and a little
delayed compared to the green beam. After I’ve de-excited my
fluorophores on the side, I’m able to look the remaining ones at
the centre. scanning technique
At the end what I have is a where I have a
smaller region but with very concentrate fluorophores.
Excitation is intrinsically a stimulation, because electrons absorb energy and
move to the next orbital.
Stimulated emission is instead a new concept: fluorescence life time (the time
the molecule stays excited) can be reduced when the molecule is shined at the
same wavelength it would emit.
This technique uses two beams:
- normal shape
- donut shape, with a hole in the centre
The excitation pulse has the same usual shape of a light beam.
Light beam is usually a Gaussian, a peak. It’s
more intense in the centre, less intense in the
sides. The intensity walking along x will start
with low intensity, than it reaches the
maximum at the centre, than it goes off on the
other side. This is a linear excitation because
the number of fluorescent molecules excited is
proportional to the intensity of light. The density, the number of molecule over
volume (for example the number of molecules in a cube of 1μm volume) that
are excited after this pulse, will have the same shape so there will be more
excited molecules in the centre of the area.
If I cut in half the intensity of the beam, the density of the molecules will be
The second beam has the opposite shape:
from zero, it goes up to the maximum, goes
down to zero in the centre and then it goes
up again to maximum and down again. This is
the quenching beam, the stimulated
If it is linear, the fraction of excited molecules
which are quenched is proportional to the
intensity, so it will de-excite more where
there are the peaks at the edge and less where it goes down in the centre.
For example: we have some flourescent molecules and we excite a fraction of
them and then with the de-excitation pulse we quench a fraction of the excited
molecules proportionally to the intensity.
If we double the intensity we also double the number of quenched molecules.
If both beams are linear we don’t gain anything because the width of the
excitation beam peak is the same of the hole in te quenching beam.
If I increase the intensity of the second beam, I increase the fraction of the
excited molecules that are quenched, and if I increase the intesity a lot I can
reach the 100% of the excited molecules quenched. But if from this intensity I
double the power, I won’t double the effect because I can’t go over 100%!
So the pattern of quenched molecules has the same pattern of light as long as
they are proportional: when I have a quench beam that de-excite 100% of the
molecule, if I augment the intensity I will shrink the hole, so I will shrink the
area where the molecules are still excited and the only fluorophores that
survive are the ones in the centre. This is non-linearity.
The blue dash is the line where the 100% of
fluorophores are quenched.
Look at the four profile patterns each one
stronger than the precedent from blue to
violet: assuming that the smallest (1) at the
maximum of the intensity de-excites 100% of
the molecule in that area, if I increase the
intensity (same wavelenght) I can quench
molecules proportionally to the intensity, and
so in a bigger area (because it’s not linear).
In this way I can shrink the area where
molecules are still excited: for each pattern the area where molecules are still
excited is smaller than the area of lower intensity pattern.
Summarize: In order to perform STED I use two beams:
Excitation beam is a pulsed laser with an
ordinary Gaussian shape
Quenching beam is a pulsed laser that is higher
in intensity and has a donut shape; the size of
the hole is usual 0,5 m.
The first pulse excites the fluorophores, the second
beam quenches them: if the STED beam has a power
zero, I can see the signal from all the fluorophores
excited. The larger the intensity of the second pulse,
the fewer are the fluorophores that remain excited.
Usually this technique allows to see the 10% of the
fluorophores, while 90% are de-excited: the 10% that
are left are the ones in the centre of the hole.
The shape of the light coming from my fluorophores is
much more narrow so I can move my resolution from
0,5µm (black spike) to 100nm (red spike).
Also you can stain your molecules with two different fluorescence proteins (GFP
and YFP) and you can excite them with the same beam and then quench them
by shooting very hard staying on the same wavelength, then looking at the
emission using a dichroic mirror to recognize the two different emission (paper
– two colours STED...).
So STED is a super-resolved optical scanning microscopy that enables 3D
By exploiting a non-linear stimulated emission, STED enables a large
improvement over the resolution limit for linear images.
Moreover by forcing the quenching of fluorophores STED minimizes bleaching
effects: the bleaching of fluorophores is proportional to the time that it stays
Total Internal Reflection microscopy (TIR)
The phenomenon is that when light goes from a
transparent material to another transparent material
and the refractive indexes are different, light that is
going through changes direction and there is
If n is larger than n , than θ has to be smaller than
1 2 1
If I go from a material with higher refraction index to one with lower refraction
index (like glass, water or oil to air) and I keep increasing θ I’ll reach the point
where exiting angle (θ ) will be 90° degrees.
2 n 2
If θ is 90°, sin θ =1 and then .
2 2 1 n 1
At this angle the exiting beam doesn’t really exit from material 1 and for angled
“exiting beam” will be completely reflected
larger than this one the : so there is
an angle above which the exiting beam doesn’t really exit and the light is
complete reflected without going outside!!
But we know that any point isn’t really a light
but has a width, so for the Snell law we
shouldn’t have light in the point of total
internal reflection in medium two but there is
a little bit of light that is just close to the
surface: if I go really close to the interface I
discover there is a little sheet of light called
evanescent wave that stays close and
doesn’t go far from the interface.
Its intensity falls just as you move away from
the surface: it starts intense and in a short
distance it goes to zero (the distance from the interface where you have this
light is far smaller than a We can use this light to illuminate something
very close to the surface.
If I want to know what are the properties of the evanescent wave, I have to
- z how intense it is
- d what’s its penetration depth (how far does it propagate?).
Both this parameters depends on the angle that light is forming with the
interface on the other side. n 2
There is the critical angle above which I have total internal
1 n 1
When this angle becomes larger intensity decreases so the best angle to have
a strong intensity is just a little higher than critical angle.
The same thing happens for penetration: if you move to larger angles,
penetration becomes narrower and narrower.
a little more the critical angle I can have the maximum of intensity and
penetration; if I keep increasing angle I will get lower intensity and penetration.
It decays exponentially.
If penetration it’s 100 nm it means that d= 100
nm; so when z is equal to d this is exponential to
the which means 1/3 of the starting intensity. So
this is the shape of the intensity moving away from the interface when the
penetration depth is 100nm.
So playing with two materials, from one with a higher
refractive index to one with a lower refractive index, you
can create this evanescence light in which the intensity
and the width depends on the angle.
But what can I do with that? I can study a cell wall because
I can illuminate just what is closer to the surface.
The refraction index of the cell is about 1.37-1.38 (1.33 is
one of water and cells have lot of other stuff), so I have to
use a material with higher refractive index (ex. glass slide
with n=1.518) and shine the light trough that.
There are two ways to do it:
External coupling on the other side I have a
beam coming that is reflecting so I can look what is
illuminated on the other side of reflection. I
illuminate the sample from below and look from the
above (accoppiamento a prisma).
Through the objective the objective stays
below: I want to shine a light on the side of the
objective so when the light comes out, it comes
with an angle tilted enough to do internal
reflection. The maximum angle you can achieve is
the one of NA, so the NA has to be larger than the
reflecting index of the cell. The best objective has
The maximum angle you can achieve is the
maximum angle you can look at and it’s basically
the angle of the NA. So comes out that NA is the refractive index (of the
material between the lense and the sample) times the sin this angle.
So basically (formula) NA has to be larger than refractive index of the
sample you are looking at (typically cell so 1.38).
7 mesi fa
Advanced optical microscopy
Introduction on the resolution of optical microscopes
Effects of the finite resolution on images
Introduction to confocal microscopy
Optical processes and techniques that can overcome the resolution limit, such as: non linear microscopy, STED,TIRF,PALM, SNOM.
introduction to nanotechnology
Nanoparticles:quantum dots, nanomag, metallic nanoparticles. General concepts, stability and bioconiugation. Optical tweezers for micro-manipulation
Microfluidics: miniaturization of lab apparatus (Lab-on-a-chip technologies), microfluidic technologies based on flux and droplets. Diffusion and mixing. Notable examples of micro-mechanicals applications.
Atomic force microscopy
Later-free optical Biosensors for the detection of not marked molecules and interactions between not marked molecules. Introduction to SPR based techniques and to methods that use guided light.
Quantitative analysis of images
Introduction to the fundamentals of computer graphics aimed to the comprehension of the informations contained in images with examples taken from microscopy, gel electrophoresis and medical diagnostic.
Colorimetry: color spectrum, Gamut, chromatic coordinates, gamma value of displays, RGB, CMYK.
Digital image types: (es. BMP,TIF,GIF,JPG)
Lossy and lossless compression
Images stacks over space (3D stacks) and time (movies). Movie file types.
Volumes and surface renderings. Tomography.
Introduction to ImageJ software as a tool to:
- extract quantitative information from an image (measurement of bands on a gel electrophoresis image, particle counting, border detection)
- Punctual, local and global filtering
- False color images applied to multi-fluorescence detection.
- Stack visualization and reslicing
- Timelapse and measurements on images over time
I contenuti di questa pagina costituiscono rielaborazioni personali del Publisher _ariiel di informazioni apprese con la frequenza delle lezioni di Advanced microscopic techniques and nanotechnology e studio autonomo di eventuali libri di riferimento in preparazione dell'esame finale o della tesi. Non devono intendersi come materiale ufficiale dell'università Milano - Unimi o del prof Bellini Tommaso.
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