Physikalische und Theoretische Chemie
Technische Universität Braunschweig
Hans-Sommer-Straße 10, 38106 Braunschweig
________________________________________________________________
Biophysical chemistry (Modul 30700)
Practical course
Single-molecule fluorescence spectroscopy
Index
1.
Keywords
-1-
2.
Introduction
-1-
3.
Theory
-1-
4.
5.
3.1. Absorption and emission of light by fluorophores
-1-
3.2. DNA Holliday Junction
-4-
3.3. Förster resonance energy transfer
-5-
3.4. Fluorescence microscope
-7-
Experiment
-8-
4.1. Preparation of a BSA/BSA-Biotin surface
-8-
4.2. Hybridization of the Holliday Junction and binding it to the surface
-8-
4.3. Surface scans and observation of single fluorescent molecules
-9-
4.4. Basic rules and safety issues regarding the experiment
-10-
Analysis
5.1. Analysis of the time traces with the Hidden-Markov model
-11-11-
6.
Time schedule
-14-
7.
Literature
-14-
Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
1. Keywords
Holliday Junction, fluorescence spectroscopy, functionality of absorption and emission spectrometer,
electromagnetic spectrum, Franck-Condon principle, Stokes shift, Beer-Lambert, extinction coefficient,
Jablonski diagram, FRET, basics of a microscope (image development, beam path ), confocal fluorescence
microscope, Hidden-Markov model (without formula)
2. Introduction
In contrast to most microscopy techniques that are limited to image acquisition, this experiment deals
with the time-resolved visualization of the dynamics of a biomolecular complex. Only the dynamics can
provide information about the function of a molecule in a biomolecular complex (e.g. DNA, protein or
lipid). In combination with known structural information of a molecule, the functionality of a molecule
can be revealed.
The aim of this experiment is to study a DNA-complex called "Holliday Junction" (HJ). To obtain this at a
single-molecular level, HJs labeled with organic dyers are immobilized on a glass surface. With a confocal
single-molecule microscope the alteration of the distances of two dye molecules linked to a HJ is
visualized by time-resolved FRET. Thus, the movement of the molecule-complex can be directly
observed.
3. Theory
3.1 Absorption and emission of light by fluorophores
Atoms and molecules can absorb light energy (photons) to change between their discreet energy levels.
By this transition not only their energetic state changes, but also their states of rotation and vibration. In
terms of their electronic structure and the yielding possible transitions and states, they absorb a various
amount of the visible (400 – 800 nm) and invisible (UV < 400 nm, IR >800 nm) spectrum - therefore they
have a specific, for their absorption characteristic color. Thereby the energy of a specific transition
always corresponds to the energy of a photon with the energy E:
𝐸 =ℎ∙𝜈 =ℎ∙
𝑐
𝜆
(1)
Here ℎ is the Planck constant, 𝜈 the frequency, 𝜆 the wavelength of the corresponding photon and 𝑐 the
speed of light. Nowadays measuring the absorption- and emission of a typical organic dye in the visible
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
and near-UV range is quite easy. So called 𝜋𝜋 ∗ ‐ transitions cause an absorption in the visible spectral
range, whereby delocalized electrons are excited to a higher energy level. The probability of the
absorption of a photon with a determined wavelength depends on the exact structure of a molecule.
Today dyes for almost every color or wavelengths of the visible spectrum are commercially available. The
shape of the absorption and emission spectra is determined by the Franck-Condon principle. The
following figure shows the absorption and emission spectra of a far red dye (ATTO655, blue =absorption,
red = fluorescence):
Figure 1: absorption and emission spectrum of ATTO655
The specific absorption A of a sample solution with the layer thickness 𝑑 can be described by the Beer-
Lambert law. It is influenced by the dye concentration in the solution 𝑐 and the so called extinction
coefficient 𝜖:
𝐴=𝜖∙𝑐∙𝑑
(2)
Here we consider only molecules, which show no chemical reaction after photo-excitation. By absorbing
a photon, a molecule is excited from the electronic ground state 𝑆0 either to the first excited singlet state
𝑆1 or the second excited singlet state 𝑆2 (figure 2, green arrows). The absorbed energy can be returned
by different processes. Directly after excitation, fast processes of vibrational relaxation and internal
conversion (figure 2, yellow arrows) occur. After some picoseconds the molecule is in a state of
relaxation, but still in a high energetic state - now it is in the energetic minimum state 𝑆1 . The following
processes are subsumed in the so called Jablonski-diagram:
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
Figure 2: Exemplary Jablonski-diagram
The fluorophore can relax to the ground state by radiative and non-radiative processes: Radiative
processes are “fluorescence” and “phosphorescence” (figure 2, red arrows). A photon is emitted during
both transitions. The difference between them is that fluorescence is a direct relaxation from the excited
state to the ground state. By intersystem crossing (isc) a change from the excited state to the triplet
state 𝑇1 is possible. In comparison to the 𝑆1 state the spin of the electron is now parallel to the spin of
the ground state. Looking at phosphorescence the triplet state is relaxed to the ground state.
Fluorescence (10-9 – 10-7 s) occurs significantly faster than phosphorescence (10-3 – 102 s). Also nonradiative transitions occur by interactions of the fluorophore with its surrounding leading to relaxations
to the ground state. Every state has a characteristic lifetime depending on the rate of radiative/nonradiative processes. The quantum efficiency of a fluorophore Φ𝑓 - the ratio of emitted to absorbed
photons - is determined by the ratio of the radiative rate to the sum of all rates depopulating the excited
state 𝑆1 . In this experiment fluorophores with high quantum efficiencies > 0.3 are used. Simultaneously
they indicate a low triplet quantum efficiency Φ𝐼𝑆𝐶 .
After several cycles of excitation and relaxation (107 − 108 times), the fluorophore is irreversible
destroyed. This effect is also known as "bleaching".
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
3.2 DNA Holliday Junction
A Holliday Junction (named after the biologist Robin Holliday) is a cross-like molecule of four single
straded DNA strands:
Figure 3: Conformations of the Holliday Junction - Left: Model of an opened HJ in absence of bivalent metal ions and salt
cations. The center of the HJ is open, the arms of the HJ point in the corners of a square. The DNA strands repel each other
because of the negatively charged backbone. Right: The HJ changes in the stacked conformation in presence of magnesiumions. This figure shows one of two possible conformations.
Such a junction is involved in the genetic recombination of two DNA-double-strands (Meiosis). In the
following figure a common model of the process of recombination of DNA is depicted:
Figure 4: Genetic recombination. a–c, Presynapsis. Spo11 (ellipses) cleaves dsDNA, yielding a covalent Spo11–DNA complex.
Endonuclease releases Spo11 bound to a short oligonucleotide, and 5' DNA strands are degraded to yield 3' ssDNA tails, which
are bound by Rad51 and Dmc1 (not shown). d–f, Crossover formation. d, Invasion of ssDNA from one end of the break forms an
asymmetric strand exchange intermediate. e, DNA synthesis (dashed line) is promoted from the invading 3' end; the second DSB
end is captured and promotes DNA synthesis. Ligation yields a pair of Holliday junctions (dHJ). f, The break apart yields a mature
product with exchanged flanking DNA. g–i, A non-crossover pathway. Strand invasion (g) and DNA synthesis (h) A transient
strand invasion complex may be dissociated, perhaps by DNA helicases, allowing new synthesized DNA to anneal to
complementary ssDNA on the other side of the break (i). Further DNA synthesis and ligation yield a mature non-crossover
product.
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
The recombination of the two DNA-double-strands depends on the one hand on the position of the HJ
which can be altered by so called "branch migration". On the other hand there are hints that it depends
on the conformation of the HJ: A vertical separation yields to a conjunction (splice). By horizontal
separation, only the inserted part (patch) is transfered to the dsDNA. Thus, the conformation of the HJ
may have influence on which genetic information is bequeathed.
These two conformations occur in the presence of metal ions (e.g. 𝑀𝑔2+ ), likewise they are present in
the cell nucleus. The HJ can switch into two stacked conformation, based on an overcompensation of the
negative charge of the DNA-backbone, by a concentration higher than 5 mM of double positive charged
magnesium-ions.
3.3 Förster resonance energy transfer
Besides the processes discussed in section 2.2, there is another process for the transition from the
excited state to the ground state. The so called FRET-transition occurs, if there are "suitable" dyes in the
immediate surroundings to each other (< 10 nm). Dyes can transfer energy radiationless via dipoledipole-coupling. Basically two dye molecules are involved in this process: A "Donor" and an "Acceptor".
The energy transfer can solely take place from the short-wavelength to the long-wavelength dye
molecule. For instance with an excitation of 532 nm the dye Cy3 (donor) can transfer its energy via FRET
to the dye Cy5 (acceptor). Cy5 emits the energy as fluorescence with a maximum around 665 nm.
Especially the distance dependency of the FRET efficiency is useful for many applications: The closer the
dye molecules are the more effective the energy transfer is. FRET sometimes is called "spectroscopic
ruler" or "optical nanometer", because with this process it is possible to resolve distances shorter 10 nm
in subnanometer resolution.
To determine the distances between the dye molecules quantitively, it is important to consider the
efficiency of the energy transfer 𝐸. If there is no energy transition, the 𝐸- value is defined as 0 and
defined as 1, if the donor-energy is completely transfered to the acceptor. The dependency of the
distance of the value 𝐸 is defined as:
𝐸(𝑟) =
1
1+�
𝑟 6
�
𝑅0
(3)
𝑅0 is the Förster-radius, at which the 𝐸-value comes to 50%. Towards a quantum mechanic calculation of
Theodor Förster 𝑅0 is defined as:
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
𝑅06 =
9 𝑙𝑛(10) 𝜅2 Φ𝐷 𝐽
128 𝜋5 𝑁𝐴 𝑛4
(4)
Φ𝐷 is the quantum yield of the donor in absence of the acceptor, 𝑛 the refractive index of the medium,
𝑁𝐴 the Avogadro constant, 𝐽 the spectral overlapping integral between the emission-spectrum of the
donor and the absorption-spectrum of the acceptor, 𝜅 2 indicates the orientation of the two fluorophoredipoles to each other. 𝜅 2 can be approximated to 2/3 by a complete free rotation of the dyes, when we
assume, the different orientation alter very fast and equitable towards each other.
To observe FRET between two dyes, an overlap between the emission-spectrum of the donor and the
absorption spectrum of the acceptor is needed. Furthermore the two dyes may not be too far from each
other - basically typical 𝑅0 -values are around 5 nm. It is possible to determine the 𝐸 -value
experimentally via the ratio of acceptor emission to the total emission of donor and acceptor:
𝐸=𝐼
𝐼𝐴
𝐴 +𝐼𝐷
(5)
Figure 5: FRET intensity in dependency of the distances between donor and acceptor (donor = green, acceptor =
red). FRET increases with lower distances between donor and acceptor
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
3.4 Fluorescence microscope
In single-molecule fluorescence microscopy, different microscopic techniques are used (confocal
microscopy, wide-field-microscopy and near-field-microscopy). In this experiment we use the confocal
setup. The parallel beam goes through the objective and is focused onto an extremely small, diffractionlimited, excitation-volume.
Figure 6: Experimental setup for a confocal measurement
In the experimental setup an excitation wavelength of 532 nm (figure 6: green lines) in combination with
appropriate detectors is used (figure 6). A dichroic mirror must be installed, so the excitation beam is
reflected into the objective and the fluorescence (figure 6: yellow and red lines) is transmitted to the
detectors. After the punctate emitted fluorescence light of the sample passes the dichroic mirror, it is
focused by the tube lens of the microscope and coupled through a pinhole. Light beyond and underneath
the focal plane of the objective can be repressed very effectively by this method and the detection
volume can be reduced to a few femtoliters. The fluorescence is split into its distinct colors by a second
dicroic mirror and then passes emmissionfilters before it is focused and detected on an avalanche
photodiode (APD). In comparison to a wide-field microscopy the confocal method does not generate an
image of the sample, but only the intensity and spectral characteristic of a spot of the detection volume
is obtained.
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
4. Experiment
4.1 Preparation of a BSA/BSA‐Biotin surface
At first the required buffer must be prepared: Dissolve one tablet of Phosphate Buffered Saline (PBS) in
200 mL water. A LabTek is used for this experiment. Purge the four cells in the middle of the Labtek three
times with 400 μL of the prepared PBS solution. Hereafter incubate the LabTek surface with a protein
layer of BSA/BSA-Biotin to bind a linker-biotin molecule, so the DNA can be attached on the glass
surface. Thus the BSA protein layer prevents interactions between the glass and the DNA. Without
isolation there would be a chemical reaction between the positive silicium atoms of the glass surface and
negative charged phosphate-back of the DNA yielding to an uncontrolled agglomeration of DNA on the
surface. Biotin is used as an anchor-molecule, which also is bound covalent on BSA. Prepare a 5:1
solution of BSA and BSA-Biotin in PBS. For this solve 5 mg BSA with 1 mg BSA/Biotin in 1.6 mL PBS - take
care to mix all prepared solutions well. For this there is a vortexer in the laboratory. After the last step of
washing purge 400 μL of the BSA/BSA-biotin solution in each of the four chambers. The solution is
incubated for two to three hours at room temperature.
4.2 Hybridization of the Holliday Junction and binding it on the surface
The hybridization takes place in a PCR tube in a volume of 100 μL. As buffer use:96 µL 1xTAE with 12.5
mM MgCl2 . For the HJ: Strand B, H, X 1.1 μL each, strand R 1 μL. This solution is heated up in a cycler to
95°C and afterwards it is cooled down for two hours to room temperature (thermo cycler program JS1).
The BSA/BSA-biotin excess in the LabTek chambers must be removed by washing three times with PBS.
Take care not to let dry the surface of the LabTek to prevent denaturation of the bound proteins. Adding
Neutravidine to the Biotin on the surface facilitates linking another biotin molecule. The affinity, so the
bond strength between Neutravidine and Biotin is one of the strongest non covalent bonds known in
biology. Each Neutravidine molecule possesses four binding-pockets. Prepare a 0.01 mg/mL Neutravidine
solution (stock solution 1 mg/mL). Per chamber 200 μL is needed. Incubate Neutravidine only in two of
the four chambers for 10 minutes. Thereafter wash each chamber three times with 400 μL PBS and then
fill them with 400 μL PBS.
In the next step the DNA can be coupled onto the Neutravidine mediated by a biotin linker that is bound
covalently to the DNA. Thereby the biotin of the DNA binds to the Neutravidine, which is strongly
coupled to the glass surface. The sample is them scanned on the confocal fluorescence microscope. The
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
following settings are used in the measuring program: Excitation: Laser 1, 1 ms/pixel integration time,
power: green 10 μW, the power is adjusted with the gauge head behind the fiber optics. Please be very
careful with the optic components, avoid any contact, since it could lead to the complete
misalignment of the setup! Before starting the scans, dilute the HJ solution in the ratio 1:1000 with PBS.
Add 1 μL of the in PBS diluted HJ sample in one of the four LabTek chambers filled with 400 µL PBS. Mix
the buffer with a pipette carefully - than start the scan as fast as possible. In the following, the Biotin of
the Holliday Junction is going to bind to the Neutravidine linked on the surface. At a certain molecule
density in the fluorescence figure (single molecules should still be differentiable), the surface is washed
twice with PBS to remove the surplus DNA-molecules (see figure 7)
Figure 7: Surface scan of single Holliday Junctions
To increase the stability of the fluorophores, we add a special composite. Therefor add 2 mM Trolox,
1% v/v GLOX and 0.1% v/v glucose in the sample chamber (ca. 750 µL total volumes). For a permanent
low concentration of oxygen in the solution during the measurement, the LabTek must be sealed. So
after mixing the buffer and the enzymatic solution close the chamber with a special silicon pad.
Afterwards scan the surface to locate the first Holliday Junctions under standard buffer conditions
(without magnesium). These "fluorescence spots", which really are single Holliday Junctions, can be
selected with the "pick and destroy" button - so the fluorescence is gathered with the time in form of
fluorescene transients. The spots can be selected with the cursor.
4.3 Surface scans and observation of single fluorescent molecules
Not only fluorescence and energy transfer between the two dyes of the Holliday Junction is can be
monitored, but also the dynamic change of confirmation of the HJ. Under certain conditions the
immobilized Holliday Junction on the surface can adopt two different conformations. This depends on
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
the magnesium concentration. The change between the two conformations occurs statistically over
time. The addition of a sufficient high concentration of magnesium causes that the HJ switches between
two stable conformations, since the magnesium can stabilize certain conformation by interactions with
the negative charged phosphate backbone of the dsDNA. A distinction of both conformations with the
time can be observed by the FRET-value, because the distances of both dyes differ from the respective
conformation significantly. Here we differentiate between a FRET population, in which both
chromophores are closer to each other (high FRET) and a population, in which they are more distant
(lower FRET). An analysis of the states based on the HMM enables to calculate the specific lifetime of
each conformations.
Now add a specific concentration of magnesium salt to the stabilizing buffer. Add 7.5 μL of the 1 M MgCl2
of the stock solution to the buffer. You get a concentration of 10 mM MgCl2. Add the oxygen removal
system and start three measurements with the LabVIEW program. You need at least 50 traces showing
FRET dynamics for a couple of seconds for a suitable statistic for each magnesium concentration. Record
more scans for MgCl2 concentrations of 25-, 50-, 100-, and 200 mM.
4.4 Basic rules and safety issues regarding the experiment
•
Do not enter the laboratories before having understood all safety instructions by the safety
delegates Angela Tiefnig (chemistry laboratory) and Dr. Guillermo Acuña (laser protection) or
their substitutes. Special clothing for protection is necessary (lab coat, gloves, protection glasses,
laser protection glasses), so make sure that you always wear them!
•
Prepare adequately for the conduction of the experiment to save you and your tutors time - it is
better to send an email to your tutor and ask what you did not understand in advance than
having to admit during the colloquium that you did not understand it.
•
Be careful with the setup! These are components of actual studies of the Tinnefeld group.
•
Act with caution; consider which chemicals are harmful and which safety measures must be
taken before you start the experiment; comply with the instructions of the assistant.
•
Single use gloves only give you limited protection! If your hands/gloves come into contact with
anything caustic/toxic you must dispose of your gloves/wash your hands immediately.
•
The used LASERs are highly energetic light sources of category 3 and therefore a danger for your
and your colleagues' eyes. Avoid your eyes to be on a level with the beam path (no chairs on
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
the same level as the laser table, cover your eyes while bending down!) and take of any
reflecting objects like watches, belt buckles, rings, neckless or similar. If anything gets broken
or does not work properly, please inform the assistant immediately - by this you save us the
error search and yourself uncomfortable questions.
5. Analysis
5.1 Analysis of the time traces with the Hidden-Markov
An essential problem analyzing single-molecule traces is the assignment of measured data (𝐸-value) to a
state, in which the molecule resides at that time. A successful approach offers the so called HiddenMarkov model. McKinney et al. applied this model successfully on FRET-traces in 2006. During this
experiment they recorded traces of single HJ's. Both donor and acceptor intensity are recorded; a time
stamp is assigned to each detected photon. To sustain a progress of intensity over time, those data must
undergo a time binning (e.g. in a 10 ms binning, i.e. all detected photons within 10 ms are added up).
This is processed with the computer program "LabVIEW".
Figure 8: Screenshot of the LabVIEW program
For that it is import that the recorded traces are cropped to the FRET fluctuations with the two black
cursors Press the “export for HaMMy?” button and set the donor channel to 1 and the acceptor channel
to 2. The binning time is 0.01 s. Export the intensity traces with the format: time - green APD - red APD
for the HaMMY-program. Take care to export the traces in the format with decimal point instead of
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
comma. The data ending must be .dat; otherwise it will not be possible to import the traces with the
HaMMy-program. Start HaMMy 4.0:
Figure 9: HaMMy 4.0 with fitted FRET traces
Chose the "traditional" method and set the number of states to 2. Now import your data from LabVIEW.
HaMMy now automatically evaluates all imported traces and saves a report-data for each trace
(figure 9).
Copy the report-data into a separate folder and open the TDP-program. Select the exposure time to
0.01 s and set the amount of traces to 2, because you can observe two transitions in this experiment
(figure 10). Import the report-data. With the button "Lock cutoffs" you can divide the screen in four
quadrants. Choose the middle of the range with the arrows above. With the arrows below the button
"pick transition", select the quadrants with the transitions 1->2 and 2->1. Select the "Fit Gaussian"
button and note the calculated parameters of the "kinetic rate" and "Stdev" (standard derivation).
Determine the parameters of the other magnesium-concentrations.
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
Figure 10: Evaluation of the reports with TDP
In the end import your data in Origin, Excel, etc. and try to fit your data points (X: MgCl2 concentration, Y:
Kinetic rates) with a monoexponential decay.
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Practical course - Single-molecule fluorescence spectroscopy - TU Braunschweig
6. Time schedule
09:00 Preparation of the BSA/biotin surface (incl. buffers)
10:00 Hybridization of HJ
10:30 Colloquium
11:15 Introduction into the confocal setup
12:00 Lunch break
13:00 Immobilization of the HJ and the first measurements
13:30 Acquisition of fluorescence transients at different magnesium concentrations
16:00 Data Analysis
18:00 End
7. Literature:
(1) McKinney, S. A.; Joo, C.; Ha, T. Biophys J 2006, 91, 1941.
(2) Vogelsang, J.; Kasper, R.; Steinhauer, C.; Person, B.; Heilemann, M.; Sauer, M.;Tinnefeld, P.
AngewChemInt Ed 2008, 47, 5465.
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