CHEM 457 Grossmont College Fluorescence Microscopy Lab 1 Report

CHEM457,CHEM457 Optical Microscopy
Youngkwang Lee
Contents
1. Brightfield Microscopy: Pixel Calibration for Measuring the Unknown
Sample Size
2. Fluorescence Microscopy 1: Determination of Diffusion Coefficients of
Microspheres by Single-Particle Tracking Analysis
• Appendix: Single Particle Tracking Analysis
3. Fluorescence Microscopy 2: Configuring a Fluorescence Filter Cube Block
4. Darkfield Microscopy: Visualizing Plasmonic Nanoparticles and Optical
Resolution
5. Characterization of Gold Nanoparticles Part I and II: Optical Properties and
Determination of Gold Nanoparticle Concentration using TEM and UV-Vis
Spectroscopy
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Brightfield Microscopy: Pixel calibration for
measuring the unknown sample size
Brightfield microscopy is the most elementary form of microscope illumination
techniques. Illumination light is transmitted through the sample and the contrast is
generated by the interaction—a combination of absorption, reflection and diffraction Figure 1. Bacteria under a
brightfield microscope
—between light and dense areas of the specimen. The name “brightfield” is derived
from the fact that the specimen is dark and contrasted by the surrounding bright
viewing field. Simple light microscopes are sometimes referred to as brightfield
microscopes.
In modern brightfield microscopy, there are two groups of planes (Figure 2).
One is conjugate field planes. The other is conjugate aperture planes. These two
planes are interdigitated each other. Once cannot visualize these two planes at the
same time as their focus is formed at physically different planes. Uniform
illumination of the sample is achieved by Kohler illumination in which the light
source is *not* focused at the plane of the specimen. As a result, the light at
specimen level is essentially grainless and
extended, and does not suffer deterioration from
Figure 2. Conjugate optical planes in microscopy. Left: Conjugate
dust and imperfections on the glass surfaces of
field planes. Right: Conjugate aperture planes.
the condenser.
To achieve Kohler illumination, the
condenser height needs to be adjusted properly.
The contrast and resolution of brightfield
microscopy also depends on the size of a
condenser aperture diaphragm. Typically, an
aperture diaphragm with the opening size of 70%
or less with respect to the back focal aperture of
an objective provides the best contrast.
In modern microscopy, digital cameras are
increasingly used as it makes it easy to do
quantitative analysis. To perform accurate
measurements, essential is understanding of how
different camera parameters impact signal
outcomes (image). The Figure 3 shows a
relationship between the image intensity and
exposure time. Exposure time is the length of
time the camera collects the signal. As shown in
the figure, the camera intensity increases as the exposure time increases. This makes sense because you detect more
photons as you let the camera collects the signal (photons) for a longer period of time. In some applications, the
intensity is quantified to extract some useful information. For example, you can measure concentration of
fluorescence molecules in a particular area by quantifying the intensity. For quantitative analysis, the intensity
should be linearly proportional to the number of photons detected. However, this is not always the case. In Figure 3,
you can find that the intensity starts off with a linear response to exposure time and then reaches a plateau. The
intensity range that shows a linear response (between B0 and I1) is called dynamic range. You should acquire image
only within the dynamic range of the camera. Exposure time greater than E1 yields nonlinear signal outputs. In a
nonlinear regime, doubling exposure time (from E1 to E2) does not double the signal. Therefore, use of this camera
setting should be avoided. The linear range of exposure time that you can use depends on gain, which controls the
amplification of the signal from the camera sensor. When a higher gain is applied, the camera responses more
sensitively—showing a greater slope of in the intensity vs exposure plot—and may reach the saturation point at a
shorter exposure time. In this lab, you will explore how different camera setting (exposure time and gain) generate
different outputs.
The objective of this lab is to 1) learn how to align/adjust a brightfield microscope, 2) determine the dynamic
range of camera, 3) understand how different camera settings (gain and exposure time) change signal outcomes, 4)
calibrate the pixel size to measure the size of unknown sample. This lab will establish basic skills and knowledge
required for using microcopy techniques with advanced contrast methods later this semester.
Experiments
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Important notice:
1) In this module, you will use one of the
microscopes shown in the section b
below.
2) Wash your hands thoroughly before
experiments. Please be careful not to
touch any lenses including objectives,
condenser, and eyepieces. You can touch
their bodies, but not lenses. If you do,
notify the instructor of it.
3) Whenever you change objectives and
load/unload a sample, lower the sample
stage. If you don’t, the objective will hit
the state and be damaged.
Figure 3. The camera intensity as a function of exposure time for
given gain.
Microscope assignment and specification:
1) Group 1 and 2: Dark-field microscope A
and B with bright-field microscopy mode
a) Model: T660C-DKO-IRIS
(AmScope), Labeled with DF A
or B.
b) Bright field microscope equipped
with dark-field microscopy function
c) Camera: CM3-U3-31S4C-CS, Color, 3.2 MP (2048 x1536), 55 FPS, SONY IMX265, FLIR
2) Group 3 and 4: Fluorescence microscope A and B with bright-field microscopy mode
a) Model: T670Q-PL-FL (AmScope), Labeled with FM A or B.
b) Bright field microscope equipped with fluorescence microscopy function
c) Camera: BFLY-U3-23S6M-C, Mono, 2.3 MP (1920 x 1200), 41 FPS, SONY IMX249, FLIR
Outline of experiments:
1) Examine the optical and operational components of the microscope
2) Open Micromanager software (camera operating software) and set the camera
3) Align the condenser
4) Acquire image using micro-manager
5) Data analysis using FIJI
A) Getting started
1) Examine labels on the microscope to recognize optical and mechanical parts
NOTE: If you use a fluorescence microscope, do not change or turn on the fluorescence microscopy
part. We use only brightfield microscopy in this module.
2) We will perform the following simple operation to examine each part of the microscope.
a. Turn on the light source for brightfield imaging: Switch on the light source (LED or tungsten
halogen lamp).
b. Control light intensity: Adjust the light intensity using the control knob. Finish with a low
intensity setting.
c. Move sample stage: Turn the x/y knob of the stage control arm.
d. Rotate focus knobs: The microscope in this course adjust focus by raising or lowering the sample
stage. Turning the coarse and fine focus knobs.
e. Rotate condenser knob: Raise and lower the condenser by turning the condenser knob.
f. Open and close the aperture diaphragm
g. Open and close the field diaphragm (iris).
h. Turn the nosepiece to change objective lenses. You must lower down the stage before moving the
nosepiece.
i. Push in and out the beam splitter that controls the light path – In: eyepiece only; out: eyepiece and
camera.
3) Finish with basic operation practice with the following setting.
– sample stage lowered by focus knob
– ample stage in the center by x/y stage moving arm
– 10x objective lens in the imaging position
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the field and aperture diaphragm almost closed
the beam splitter in the “out” position.
LED or lamp intensity set to minimum.
Those who use a fluorescence microscope for a bright field microscopy lab,
NOTE: If you accidently changed the fluorescence microscopy part settings, change them
back to the following settings.
o epi shutter closed
o filter cube position in the center (in position O)
B) Camera setting
1) First, log in computer.
a. ID: COS-SMBClabcsl222
b. PW: ChemSMBCcsl222@
2) Open Micro-manager on desktop. Click ok.
3) Set the configuration setting and image settings as shown in the figure 4.
a. Exposure time 10 ms. Exposure time is the length of time the camera collects the signal. If you set
a greater value, camera collect signal for a longer period of time. Consequently, the intensity will
increase.
b. Temperature is reading-only value.
Figure 4. The main control window of Micromanager.
4) Start camera by clicking the Live icon located on the left side. This will display the sample image in real
time.
5) You will see the two popup windows — Preview that shows the live image and Inspect that shows an
intensity histogram of the selected image.
a. Check off Autostretch. This function automatically adjust contrast.
6) Now you completed a basic camera setting.
C) Align brightfield condenser.
1) To align a condenser, we will visualize a field diaphragm, which is on focus when the condenser is
properly aligned. Noe that we *cannot* align the condenser by directly visualizing the condenser aperture
because it is not focused on the sample plane, which a camera visualizes. Recall that they are two groups of
conjugate planes in optical microscopy—(1) conjugated aperture planes and (2) conjugated field planes.
They are interdigitated and not visualized at the same time.
2) Adjust the opening size of the field diaphragm such that the bright circle spans about half of the image.
3) Focus the sample by turning the focus knob. Use a 10x objective. You should see a sample image as shown
in Figure 5.
a. The intensity histogram in the inspection window should *not* saturate. See the figure 7 for an
example of a saturated image. Target the max intensity value of the histogram in a range between
20% by lowering LED power. (see the pre-lab video, if this is not clear.)
4) When the condenser is well aligned (at the right height and centered), you should be able to see the sharp
edge (blades) of the field diaphragm.
a. If not, you need to align its height (z) and x-y position.
b. Align z (vertical) position of the condenser until you see the sharp edge feature of the field
diaphragm. The well-aligned example is shown in C in Figure 5.
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Figure 5. Condenser alignment.
c. If your condenser is off in x/y direction, center it using two centering knobs in the condenser (the
long two knobs at an angle).
5) Adjust the condenser aperture diaphragm.
a. The condenser aperture diaphragm controls the amount and angle of illumination light. The best
contrast is typically obtained when the opening size of the condenser aperture is about 70% or less
of the back focal aperture of the objective.
b. Do the following steps to adjust the condenser aperture.
• Remove one eyepiece.
• Look down through the empty eyepiece path on the microscope.
• You will see both the aperture diaphragm (adjustable hole) and the back focal aperture of
the objective.
• Open and close to see how it change.
• Adjust the aperture diaphragm to fill about 70% of the back focal aperture of the
objective.
• Replace the eyepiece.
6) Take aligned condenser images Lab Report Figure 1
1) You need to submit an aligned condenser image that display centered and sharp blades.
2) Take a snapshot by clicking Album (recommended) or SNAP.
a. Sometimes SNAP creates an error. If it happens, try again with Album.
b. Stop Live, if camera is on.
c. Save image by clicking the disk icon in the right bottom corner of the preview. Enter
appropriate file name so that you can find the image later (enter magnitude information).
D) Understanding parameters controlling image quality.
1) Sometimes the image appears with poor contrast. There are various reasons for this. A key to highquality imaging is to make balance between the light source intensity and camera sensitivity (or
exposure time). There is more than one condition that produces a high-quality image.
2) We will play with several parameters to understand how those impact the image quality. Yellow
indicates data acquisition. Blue indicates the data analysis after the lab.
a. Let’s first discuss what we want to avoid. Sometimes you will find your image is too bright (bad)
as shown in figure 7. This typically creates a white out image. You will also find that the intensity
histogram in the inspect window is saturated as sown in figure 6—essentially there is no contrast!
b. To avoid this, the highest intensity of the histogram should be about 10% of the max intensity (For
a 16-bit camera, it is 65,535. Therefore about 6,000 is a good target.)
c. The contrast needs to be adjusted properly. Adjusting contrast here does *not* mean changing
absolute intensity values. It changes the way to display the image. You can check off autostretch or
adjust manually in the intensity histogram by defining the lower and higher ends of the diagonal
line (indicated with triangles).
d. Adjust the size of the field diaphragm opening to about half of the field of view as shown in
Figure 5.
e. LED light source: Increase and decrease the light source power and observe how it changes the
intensity histogram in the inspection window. You can also display the intensity of the particular
position of the image. To do this, hover your mouse over the area of image you are interested in.
The intensity should be displayed in the main menu bar of the Micromanager.
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The intensity
reached the max
value (saturated)
Figure 7. Saturated white out
image
(Bad, too bright imaging
condition).
Figure 6. Inspection window showing the intensity histogram (black graph).
i.
ii.
f.
g.
h.
i.
j.
k.
NOTE: Note that although the actual intensity changes (as shown in the histogram), your image
may remain the same. That is because you have autostretch (autocontrast) is on.
Finish the LED test with the highest intensity in the histogram at 10%. If you cannot get this range,
move to the next step.
Turn on autostretch.
Condenser aperture diaphragm: Lab Report Discussion d Change the size of opening and
observe how it changes the intensity histogram. Does it change the visualized area (white circle)?
Change back the condenser aperture diaphragm setting to have the highest intensity in the
histogram at 10% (Some microscopes may not be able to get the highest intensity to 10%. 20% is
also ok.)
NOTE: Again, your image may remain the same although the intensity histogram changes, if
autostretch is on.
Field diaphragm: Lab Report Discussion c Change the size of opening and observe how it
changes the intensity histogram.? Does it change the visualized area? Does it change the intensity
of the white background area? Qualitatively describe your observation in the lab report
Exposure time: Lab Report Figure 4 Exposure time is the length of time the camera collects the
signal. Acquire images by clicking Album (IMPORTANT!) with different exposure time at 10,
20, 40, 60, 80, and 100 ms. If you use Ablum, the images will be stored as a single stack file
(multiframe image) in the order of acquisition. For example, the first frame is 10 ms, the second
frame is 20 ms, and so on. Do not save it as multiple files!
Analysis is individual work and should not be shared: See “The image analysis protocols, 2)
Measurement of the Average Intensity in Region of Interest (ROI)” at the end of the lab
manual. In your lab report draw a plot of the mean intensity of the center area of images as a
function of exposure time. Determine and visually indicate the dynamic range of a camera on the
y-axis. The dynamic range is a range of the intensity that shows a linear response to a parameter
being controlled (exposure time in this measurement).
Gain: Lab Report Figure 5 Gain is a digital camera setting that controls the amplification of the
signal from the camera sensor. The higher gain, the higher signal. But the gain may or may not
increase a signal to noise ratio as an electronic noise is also amplified. Set an exposure time to 10
ms. Ensure the highest intensity in the histogram is about 10%. Acquire images with different gain
at 0, 1, 2, 4, 6, 8, 10, 15, 20. In your lab report, determine and visually indicate the dynamic range
of a camera on the y-axis. The dynamic range is a range of the intensity that shows linear response
to a parameter being controlled (gain in this measurement).
Lab Report Figure 6 Now we will examine how different combinations of gain and exposure
time set the camera sensitivity. Acquire images with a gain of 1 while changing exposure time at
10, 20, 30, and 40 ms. Make the same measurements (10, 20, 30, and 40 ms exposure) with a gain
of 3 and 6.
In your lab report (*not* during the lab), draw plots of intensity as a function of exposure time.
Plot data obtained with different gain values into the same graph (only one graph!).
E) Take images of the calibration standard Lab Report Figure 2
1) Fully open the field diaphragm
2) Take one image at each magnification (10x and 40x).
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3) Save image by clicking the disk icon in the right bottom corner of the preview. Enter appropriate file name
so that you can find the image later (enter magnitude information).
4) To change the objective, slightly lower the stage using the focus knob to avoid crashing the objective lens.
5) Carefully turn the nosepiece to change the objective to a higher magnification.
6) Readjust the focus and other settings if necessary. Do NOT change the x/y/z position of the condenser.
7) Take images.
F) Finish the measurement
a. Lower the stage
b. Lower the condenser
c. Change the objective lens to 10x magnification
d. Turn off the light source
e. Place the sample back to the container
f. Close micro-manager software
——————————–After this line, it is individual work. Do not discuss each other—————————–Image Analysis Protocols for Brightfield Microscopy
1) Determine the size of unknown samples
In order to determine the size of unknown sample, we will first have to calibrate the pixel dimension of the
microscopy system. The calibrated pixel dimension is in a unit of µm/pixel. Then you will need to calculate the size
of unknown using the following equation:
The length of sample (µm) = the number of pixels spanning sample (pixels) x calibrated pixel dimension (µm/pixel).
The following procedure shows how to do pixel calibration using a standard calibration sample and determine the
sample size.
Calibration of the pixel dimension
This is to calibrate the pixel size for the microscopy system with a given objective lens and camera.
1. Open a calibration standard image obtained with a given objective in FIJI.
NOTE: You need to calibrate the pixel dimension for all magnifications at which the images are acquired.
2. Adjust the contrast if necessary
a. Image > Adjust > Brightness/Contrast, or shortcut Ctrl+Shift+p
3. Set image properties to the unit pixel
a. Image > Properties
b. Set unit of length: pixel
c. Set Pixel width: 1
d. Set Pixel height: 1
e. Set Voxel depth: 1
f. Other settings do not matter.
g. Click OK.
4. Zoom in if necessary.
a. Click the magnification icon !
.
b. The right click over the image will zoom in. The left click will zoom out.
5. Measure the distance between the calibration lines in pixel.
a. Click the line measurement icon !
.
b. Draw the line connecting two points between which you want to measure the distance.
c. Measure the distance in pixel.
i. Analyze > Measure, or Ctrl + M
ii. Read a value for Length. This value is in pixel.
d. Perform two more measurements (Step 5.b-c), then take the average value of all three
measurements.
6. Set scale.
a. Now you are going to enter the calibration information to FIJI.
b. Analyze > Set Scale
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c. Enter the average pixel distance to Distance in Pixel.
d. Enter the known distance. See the calibration standard drawing for the distance between lines.
Drawing of the Calibration slide
e. Enter the unit.
f. Click ok.
7. Confirm the calibrated pixel scale.
a. Image > Properties
b. Screen capture the Image Properties window for the report.
2) Measurement of the Average Intensity in Region of Interest (ROI)
1. Adjust image contras if needed. Image > Adjust > Brightness/Contrast. Click Auto. Do not click Set or
Reset.
2. We will first define ROI. Click the circular area button shown below.
3. !
4. Draw a circle within the white illuminated area at the center.
Note: You can make perfect circle by holding the Shift key. You need to make roughly the same size or circle
for the remaining analysis. So, record the diameter of ROI, which is displayed in the main ImageJ window
(w=xxx, h=xxx).
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5. If you have a stack image (multiframe image; you will get an image stack if you acquired the image by
Album during the lab), we can automatically measure the average intensities of all frames in stack file.
6. Go to Image > Stacks> Plot Z-axis Project
Note: You have to be on the first frame of the stack image when you execute Plot-Z-axis Project.
7. Intensity plots will show up. The x-axis label is often not correct. It is the mean intensity (y) as a function
of frame (x).
8. Click List at the left bottom of the graph to get text data. The first column is frame number. The y axis is
the mean intensity.
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How to prepare lab reports of the optical microscopy labs (Apply to all lab reports)
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All lab reports will be submitted via Canvas.
Lab reports include questions regarding the lab and theory, results (data presentation) and discussion.
Figures in all lab reports should have all necessary information (e.g. axis labels, legends, etc.). Do NOT
put the figure title on the top of the figure.
An example of scientific figure presentation is shown below.
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All TIF files must be changed to JPEG. You can change the microscope image format from Tiff to JPEG in
FIJI (File > Save as).
A table should have a caption.
The easiest way is to prepare figures in ppt and save it as a figure. See this page. https://
www.howtogeek.com/403177/how-to-save-powerpoint-objects-as-pictures/
General formatting information:
o Font: Arial or Times New Roman
o Font size: 10 for arial, 11 for times new roman
o Line spacing: 1
o Upload file format: pdf
Chem 457 Brightfield Microscopy Lab Report
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SECTION:_____, GROUP:_____, NAME:______, Red ID_______
1) Results
Present the following images and plots. All data should be displayed as a figure with a proper
format as shown in the previous page. The data (Figure 4-6) needs to be analyzed as described in
the lab manual. Below are not figure caption. You need to write appropriate figure caption that
explain the figures.
Figure 1. displaying aligned condenser image
Figure 2. displaying a standard sample at each magnification
Figure 3. displaying a screen capture of image properties windows of FIJI with calibrated pixel
information.
See the image analysis protocols “Calibration of the pixel dimension”
For Figure 4-5, you may or not reach a saturation point. Either is fine.
Figure 4. plot displaying the intensities at different exposure times.
See “The image analysis protocols, 2) Measurement of the Average Intensity in Region of
Interest (ROI)”
Figure 5. plot displaying the intensities at different gain values
Figure 6. plot displaying the intensities with different combinations of exposure times and gain
values.
2) Discussion
a. Briefly explain how condenser can be aligned based on the concept of optically conjugated plane
groups. (Do not discuss the operation procedures).
b. Determine the diameter of microemulsions (download from Canvas). Measure at least 10
emulsions. Report average and standard deviation. The image is acquired with a 10x objective.
Indicate if you used fluorescence microscopy or darkfield microscopy in the brightfield lab.
See the image analysis protocols “1) Determine the size of unknown samples”
c. How do the field of view and intensity change when you close and open the field diaphragm?
Explain your observation based on theory of optical microscopy discussed in the class.
d. How do the field of view and intensity change when you close and open the aperture diaphragm?
Explain your observation based on theory of optical microscopy discussed in the class.
e. Discuss Figure 4-5. You may explain the results, trends, and any findings (how exposure time and
gain contribute to outcome of images and intensities). You do not need to display any images, but
need to explain the plots and images. See the canvas page of brightfield microscopy lab for
introduction to signal, gain, and dynamic range.
f. Discuss Figure 6. You may explain the results, trends, and any findings.
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Fluorescence Microscopy 1. Determination of
diffusion coefficients of fluorescence microspheres
by single-particle tracking analysis
Single-particle/molecule tracking is a powerful technique for
measurements of trajectories of individual proteins or lipids in
Figure 1. Single molecule tracking of
living cells. Molecular interactions and chemical reactions often
individual proteins.
lead to change in diffusion behavior of molecules. Thus, analysis
of diffusion dynamics allows us to study biochemical reactions as
they happen, inside the living organism, one molecule at a time.
Scientists found that the molecules in cells repeatedly undergo
cycles of fast-diffusing monomers and slowly diffusing or
immobile, transient molecular complexes greater than dimers.
These observations established foundational knowledge about
mechanisms and dynamics of biochemical reactions in cells, such
as those involved cell signaling and gene regulation.
These experiments largely rely on estimation of diffusion coefficients of single objects such as a particle or a
fluorophore that is conjugated to the molecule of interest. The objective of this lab is to determine diffusion
coefficients via an experimental method that is widely used in modern biophysics—single particle tracking. We will
further extend this experimental method to the determination of an important chemical constant such as Avogadro’s
number.
The diffusion coefficient D of spherical particles undergoing Brownian Motion in a quiescent fluid at uniform
temperature is described by the Stokes-Einstein equation:
D=
RT
NA6π ηa
Where R is the gas constant, T is the temperature, NA is the Avogadro’s number, η is the dynamic viscosity of a
medium, and a is the radius of the spherical particle (or hydrodynamic radius of molecule). As described in the
equation, the rate of diffusion increase as the temperature increases. As the solvent viscosity and the radius of the
particle increase, the particle diffusion slows down. This principle can be applied to estimate diffusion coefficient of
molecules. The formula is historically important since it was used to make the first absolute measurement of
Avogadro’s number so confirming molecular theory.
The probability density for a particle to move a distance r in time interval t when the particle can be ascribed
diffusion coefficient D is
r
r2
ρ(r, t, D) =
exp −
.
2Dt
( 4Dt )
The equation can be used to experimentally determine the diffusion coefficient by fitting the step size
distribution obtained from single-particle tracking experiments. In this lab, we will record movement of different
sizes of microspheres using a fluorescence microscope. Then individual diffusion trajectories will be analyzed to
construct step size distribution.
Experimental Procedures
Microscope specifications
• Model: T670Q-PL-FL (AmScope)
• Infinity-corrected optical system
• Lens: 10x NA0.25 FLOR, 40x NA0.65 FLOUR, 40x NA0.65 PLAN
• Condenser: NA1.25 with iris diaphragm & filter
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Transmitted illumination: Köehler, LED
Fluorescence light source: mercury-vapor arc-lamp, 100W
Filter set B: excitation 490 nm (band width not specified); dichroic 510 nm; emission 530 nm (band widthnot
specified)
Filter set G: excitation 545 nm (band width not specified); dichroic 580 nm; emission 590 nm (band widthnot
specified)
Camera: Blackfly 2.3 MP (1920×1200) mono USB3 vision, Sony Pregius IMX249, FLIR
Caution & important notice
1) When a fluorescence lamp is on, a mercury lamp housing on the back of the microscope is extremely hot. Do
not touch a lamp housing. You will be burned.
2) Wash your hands thoroughly before experiments. Please be careful not to touch any lenses including
objectives, condenser, and eyepieces. You can touch their bodies, but not lenses. If you do, notify the
instructor of it.
3) Whenever you change objectives and load/unload a sample, lower the sample stage. If you don’t, the
objective will hit the state and be damaged.
4) Each group member must acquire stream images at least one sample. If your group already acquired all
data before your turn, acquire your own data additionally. You are welcomed to make extra measurements
if you have enough time.
Experimental overview
1) Examine microscopy labels and parts
2) Bright field imaging and condenser alignment
3) Fluorescent microbead sample preparation
4) Fluorescent microbeads imaging
A. Getting started: Examine microscope parts and get started
1) Before loading the sample, take a moment to get familiarized with the microscope parts.
• Examine labels on the microscope.
• LED light source for brightfield imaging: Switch on the LED light. The switch is located at
theback. Adjust the LED intensity. Finish with a low intensity setting.
• Turn on a mercury arc lamp for fluorescence imaging: If the mercury lamp is not on, switch
on.NOTE: Never turn off. It must be cooled down for an hour to turn on again.
• Sample stage: Move the sample stage in x-y directions using the stage-moving arm.
• Focus knobs: Raise and lower the sample stage by turning the coarse and fine focus knobs.
• Condenser knob: Raise and lower the condenser by turning the condenser knob.
• Open and close the aperture and field diaphragm. Leave them open.
• Turning the nosepiece to change objective lenses. You must lower down the stage.
• Push in and out the filter cube block.
•
Push in and out the beam splitter that controls the light path – In: eyepiece only; out: eyepiece
andcamera. Finish with “out”.
• Push in and out the epi shutter.
• Finish with basic operation practice with the following setting.
– epi shutter closed
– filter cube position in the center (in position O)
– sample stage lowered
– sample stage in the center
– 10x objective lens in the imaging position
– the field and aperture iris fully opened
– the beam splitter in the “out” position.
– LED intensity set to the lowest line
B. Sample preparation
1) NOTE: Each group prepare one slide and share the sample. All students should participate in the
sample prep.
2) You will image fluorescent microspheres and analyze diffusion dynamics of individual particles. The
table below is information of the fluorescence microspheres used in this experiment.
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Partic
le
ID
Diamet
er
Fluorescence
(excitation, emission
(nm))
Core/Surface
functionalizati
on
Filter
set
Diluti
on
A
1 µm
Suncoast Yellow (540,
600)
Polystyrene/COOH
G
1/300x
B
2 µm
Suncoast Yellow (540,
600)
Polystyrene/COOH
G
1/150x
C
5 µm
Dragon Green (480, 520)
Polystyrene/COOH
B
1/50x
Table 1. Information of fluorescence microspheres.
3) Prepare imaging chamber with a three hole-punched double side-sticky tape attached onto the
slideglass.
4) Resuspend particles. Set micropipette to 20 uL. Vigorously pipette up and down stock solution
multipletimes to homogeneously mix particles. It is important to make sure that particles are well
separated one another.
5) Dilute particles. Add 1 µL of particle solution in appropriate volume of deionized water (DIW). Use
thedilution factors in Table 1. For example, to yield 1/50x dilution add 1 µL of particle #1 into 50 µL of
DIW. The recommended dilution factor will yield a proper particle density for single particle tracking.
But you may need to adjust the dilution factor as needed. It depends on your pipette accuracy.
6) Briefly sonicate the solution for 30 sec to thoroughly mix diluted solution. Since the particles tends to
aggregate, the sonication is important to achieve well-dispersed particles. NOTE: You need to mix well
every time you use it. If your particles are stationary when you image, you need to make the
sampleagain.
7) Drop 10 µL of each diluted particle solution onto the center of the hole.
8) Close the chamber with a glass coverslip and gently press the periphery of the imaging chamber using
aplastic tip. NOTE: Do not apply too much force or press the imaging window. It will be broken.
C. Open image acquisition software (Micro-manager) and set the camera
1) Launch the microscope control software Micro-manager on the desktop. Click OK.
2) Configure camera settings as shown in Figure 2.
i) Exposure 10 ms
ii) Gain 0
iii) PixelType 16 bit
iv) Temp is a reading only value.
v) You may not see the other settings. It is ok.
!
Figure 2. Camera settings. Exposure time: the length of time to collect signal. Gain: a degree of signal
amplification. Bit: image depth.
D. Brightfield microscopy imaging Lab Report Figure 1
1) NOTE: the operation instruction for the brightfield microscopy is brief since you already finishedthe
bright field microscopy lab module.
!14
2)
3)
4)
5)
6)
7)
8)
9)
CHEM457,
We will first visualize the particle C (5 µm) and alight the condenser.
Position the sample C under the 40x objective.
Live the camera.
Focus the sample.
Tips: You may need to adjust aperture diaphragm and light intensity to have proper illumination
condition.
Condenser adjustment. Adjust the condenser height and x/y position until you see the sharp
fielddiaphragm image in the center.
Tips: You need to partially close the field diaphragm to see the edge of the blades.
Once the condenser is aligned, open the field diaphragm all the way.
Readjust aperture diaphragm and LED power, if necessary.
To reduce data size, we will use only the center area of the CMOS camera sensor. So we will
defineRegion of Interest (ROI).
i) Snap or Album to take an image.
ii) Click the middle ROI button ”
. It will set the ROI to the center quarter of the chip.
iii) Click Snap or Album to accept the ROI. Now you see that we are using only the center quarter of
thechip.
10) Take an image by clicking Snap or Album (if you see an error message in Snap).
i) In contrast to Snap, Album does not stop camera. So you have to click Stop Live.
After save image, you have to close the Album window. Otherwise the image will be overwritten.
NOTE: You need to save image files and record imaging conditions (e.g. exposure time, gain,
magnification, etc.) to prepare the lab report.
E. Fluorescence microscopy imaging Lab Report Figure 1
1) Now we change the microscope setting for fluorescence imaging.
i) Stop Live if your camera is still in Live.
ii) Lower stage.
iii) Locate the sample C under the 40x objective.
iv) Turn off the LED light. The switch is on the back of the scope.
2) Slide the fluorescence filter cube block left or right to position an appropriate filer unit.
i) See Table 1 to find the correct filter cube for each particle. Ask a TA or Dr. Lee if you are not sure
about it
3) Open the epi fluorescence shutter. You will see the excitation light coming out of the objective.
4) Set exposure time to 10 ms.
5) Start Live.
6) Raise the stage with the coarse focus knob to focus. You should be able to see the diffusing particles
onthe screen.
7) Fine adjust the focus with the fine focus knob.
8) Adjust the exposure time, if necessary.
i) Make sure the camera is NOT saturated. When saturated, the particles appear just white dots,
ratherthan spheres with gradual contrast.
9) Stop Live.
10) Take an image by clicking Snap or Album (if you see an error message in Snap).
i) In contrast to Snap, Album does not stop camera. So you have to click Stop Live.
ii) After save image, you have to close the Album window. Otherwise the image will be
overwritten.NOTE: You need to save image files and record imaging conditions (e.g. exposure time,
gain, magnification, etc.) to prepare the lab report.
F. Record movie of diffusive particles Raw Data for Lab Report Figure 2 and Table 1
1) Now we will search for a good area – a large number of particles that are well separated one another.
Avoid particles that interact with each other. You should have at least 20 particles in the field of view.
2) You will acquire the movie only with the 40x objective.
3) Acquire Movie: Click Multi-D. Acq ”
i) Check box Time points & Save images.
ii) Set the following parameter.
(1) Number: 200 (Total frame)
(2) Time interval: 0.2 s
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.
CHEM457,
(3) Exposure time: 10 ms or the exposure time you optimized in the snap acquisition.
(4) Define a save directory root. The save directory is on the desktop. Make a directory with
yoursection and group number. For example, 9 am Group 1. You can make more sub
directories as needed.
(5) Check Save as a stack image.
(6) Define the file name.
(a) A typical file name would follow the following format:
Sample info_XX frames_exp XXms_interval XXms_XXlens
Example: Fluorescence_100 frames_exp 10ms_interval 200ms_10xlens
iii) Click Acquire to start recording.
iv) Play the acquired movie. NOTE: Check the total number of frame and error frames.
G. Acquire brightfield images, fluorescence images, and movie for the sample B and C. Lab Report Figure 1
Note: in this module, you need to acquire the following
data.For each sample (A, B, and C):
• One brightfield images with 40x objectives.
• One fluorescence images with 40x objectives.
• One florescence movie (200 frame long).
H. We will calculate diffusion coefficient, D, from single particle tracking data. Lab Report Figure 2 and Table 1
i) The detailed analysis procedures were discussed in the class. The written protocols for single
particletracking and analysis templates are uploaded in Canvas.
ii) Analyze step sizes of particle diffusion using TrackMate in ImageJ.
(1) USE the calibration pixel dimension uploaded in the Canvas for the 40x
objective(a) You must enter the calibrated pixel dimension in Image Properties of
ImageJ.
(2) Refer to the practice lab manual and video if needed. You can use the template
spreadsheetprovided for the practice lab.
(a) To see these documents, go to Canvas Module “Practice: Estimate Diffusion
Coefficientsby Single Particle Tracking”.
(b) Note that the pixel dimension used in the practice lab is different from this
“Fluorescence microscopy lab #1”. You need to accordingly adjust the
analysisparameters. See the practice lab manual for more detail.
iii) Freely diffusing particles undergo random Brownian motion. Diffusion coefficient, D, can
becalculated by fitting the step size distribution with simple Brownian diffusion model.
iv) The probability density for a particle to move a distance r in time interval t when the particle can be
ascribed with diffusion constant D is 𝑝(𝑟, 𝑡, 𝐷) =
2
𝑟
2𝐷𝑡
𝑟
exp (−
). By fitting the step size
4𝐷𝑡
distribution to this equation, the diffusion coefficient D (µm2/s) can be calculated. Note that the p(r, t,
D) is the probability density, not probability.
!16
Chem 457 Fluorescence Microscopy Lab 1 Report
CHEM457,
SECTION:_____, GROUP:_____, NAME:______
Title: Determination of diffusion coefficients of fluorescence microspheres by single-particle
tracking analysis
1.
Results You need to present the following data. Provide appropriate figure and table caption.
a. Figure 1. Brightfield and fluorescence images of A, B, and C acquired with 40x lens.
(No need to display 10x images).
i. Add a scale bar.
ii. See this page for how to add scale bar: https://www.swarthmore.edu/NatSci/
nkaplin1/scalebar.htm
iii. Note: The pixel calibration information is on the Canvas page of FM1. As we
already know the calibrated pixel dimension, we do not need to do step 1-3. You
can directly set the scale starting with step 4.
iv. A scale bar example is shown below
!
b. Figure 2. Step size distribution of particle A, B, and C with fit curves. All plots and fit
curves should be plotted in the same graph. Place all your data into a single graph so
that we can directly compare three particles.
c. Table 1. Estimated diffusion coefficients for particle A, B, and C. This table should
include particle diameter information.
d. Figure 3. Plot of experimentally determined diffusion coefficients and theoretical values
as a function of particle diameter. (See the introduction of the lab for more information).
Constants are provided in Discussion b below)
2. Discussion
a. Discuss a trend of estimated diffusion coefficients. Compare it with theoretical prediction.
Any noticeable differences? Discuss about the potential source of deviation (you will
receive the full mark for this discussion as long as your answer scientifically sounds
right). (Discussion of Figure 3)
b. Determine the absolute Avogadro’s Number using an estimated diffusion coefficient of 1
µm particle. Temperature is 23 °C. The dynamic viscosity of water at 23 C is 0.9321
mPa.S. (See the introduction of the lab for more information)
c. Suppose you are determining the diffusion coefficient of the following molecule in a
homogeneous liquid like water. List molecules in the order of higher diffusion coefficients
assuming they are in the same solution. Provide justification. (Tip: What would be the
most critical factor that determine the diffusion coefficient of these molecules in the same
liquid?)
i. Molecules: H2, vitamin-c, 20-bp-long single strand DNA, glucose oxidase protein
!17
CHEM457,
Fluorescence Microscopy 2. Configuring a
fluorescence microscopy filter cube
A fluorescence microscope is an optical microscope that uses fluorescence emission as an imaging contrast
method. It is one of the most widely used microscopy technique in research because of its excellent sensitivity and
chemical selectivity. In fluorescence microscopy the specimen is illuminated with light of a specific wavelength
which is absorbed by fluorophores, causing them to emit light of longer wavelengths. Then the fluorescence
image is visualized via a camera. To achieve a high-quality fluorescence image, it is essential to efficiently excite
fluorophores, collect emitted photons, remove the
excitation light that is reflected from the substrate. The
Figure 1. The anatomy of a fluorescence filter cube.
optical component that are used for this purpose is a
fluorescence filter cube (block). A fluorescence filter
cube (block) directs beam paths and selects appropriate
wavelengths in fluorescence microscopy. Configuring
appropriate filter sets according to the spectral features
of a fluorophore has a great impact on the performance
in fluorescence microscopy. There are three categories of
filters to be sorted out: excitation (exciter) filters,
emission (barrier) filters and dichromatic beamsplitters
(dichroic mirrors) that are usually combined to produce a
filter cube similar to the one illustrated in Figure 1.
Proper selection of filters is a key to successful
fluorescence microscopy experiments.
Excitation (exciter) filters permit only selected
wavelengths from the illuminator to pass through on the
way toward the specimen. Emission (barrier) filters are
filters which are designed to suppress or block (absorb) the excitation
wavelengths and permit only selected emission wavelengths to pass
toward the eye or other detector. Dichromatic beamsplitters (dichroic
mirrors) are specialized filters which are designed to efficiently reflect
excitation wavelengths and pass emission wavelengths. They are used
in reflected light fluorescence illuminators and are positioned in the
light path after the exciter filter but before the barrier filter.
Dichromatic beamsplitters are oriented at a 45 degree angle to the light
passing through the excitation filter and at a 45 degree angle to the
barrier filter as illustrated in Figure 1.
Figure 2. Fluorescence filters.
Performance of a set of excitation and emission filters for a given fluorophore can be quantitatively compared
using the equation shown below:
The equation estimates the degree of spectral overlap between the excitation filter and the excitation spectrum of a
fluorophore and between the emission filter and the emission spectrum, which determines the efficiency of
excitation and collection of fluorescence emission.
The objective of this lab is to select appropriate excitation and emission filters for a given fluorophore. To do
this we will measure the transmission efficiency of fluorescence filters (Figure 2) and compute the degree of
spectral overlapping between optical filters, absorption and emission spectra to determine the effective brightness.
This knowledge will allow you to better understand the signal detection mechanisms and to optimize/evaluate
filter sets in a fluorescence microscope.
Experiments
Measurement of Transmission Efficiency of Fluorescence Filters and Data Analysis
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CHEM457,
1) Turn on the computer and software.
a) Log into the computer with the student account. Ask the password to the instructor.
b) Open Cary 50 UV/Vis Spectrophotometer software by clicking the Scan icon on the desktop. You can
also find the icon on the start menu of Windows. Wait until the high-frequency sound stops and make
sure that the software shows online on its top status bar.
2) Setting up measurement conditions
a) Click Setup on the left top to open setup window. Set up Start and Stop wavelength, Y mode, Beam
Mode, Scan controls, and Display Options as shown in the Figure 3. The detailed setting info is in the
figure caption.
b) Click Baseline tap. Select Baseline correction. Choosing this option will ask measurement of a blank
sample to acquire a background signal before measuring samples of interest.
c) Click Auto Store tap. Select Storage on [prompt at start].
3) Sample measurement
a) Measure a blank sample by clicking Baseline on the left side of the main screen. No sample is loaded
on the sample holder for the blank measurement.
b) Attach excitation or emission filters on the sample holder. You will find double side stick tape
attached on the right side of the holder. If
not, talk to the instructor.
c) C l i c k S t a r t . E n t e r f i l e n a m e – Figure 3. setup window of UV/vis spectrometer.
SectionX_GroupX_filter transmission. Start 800; Stop 400; Y min -5; Y max 105; Beam mode: Single
Write sample name – Filter #X. Unload Beam; Scan Control: Slow; Display Option: Overlay Data.
sample, and then load next sample. Enter
sample name. Repeat the above
procedure to measure the all other filters.
Click finish. You will be given 6 filters in
total (1,3,4,5,6, and 7. Note that we do
NOT have filter #2).
4) Export data
a) C l i c k f i l e – > S a v e D a t a A s – >
Spreadsheet Ascii (*.CSV) -> enter name
as Filter transmission spectra.
5) Shutdown the spectrophotometer
a) Close the software and logoff the
computer. Do not turn off the computer.
6) Data analysis
a) You can analyze and plot your data using
any of software that you are comfortable with and accessible to. There are a number of scientific data
analysis software including Excel, Origin, Igor, Matlab, Prism, and more. If you are familiar with
computer languages you can use free data analysis platforms such as Anaconda, which is based on
Python. Here, general analysis processes using Excel is described below. But feel free to use whatever
software as long as you can do the same analysis.
b) Open the saved ascii file in Excel. Copy and paste the entire spectrum data into analysis template
file. Paste data into orange colored cells. File: FM2_Analysis template_Filter transmission
spectra.xlsx
c) You will see transmission values for each spectrum as a function of wavelength. Wavelength values
include the fractional part. To make analysis easy, we will convert wavelength values to integers
(this will decrease a resolution of a spectrum, but it is fine for our purpose.). The converted
wavelength is shown in column A (blue). Check the ROUND function and get familiarized with it.
You can see a function by clicking one of the values.
d) Next we will get normalized transmission efficiency for each wavelength (integer). For this, we use
the following function =INDEX(C$5:C$805,MATCH($O5,$A$5:$A$805,0))/100. This is an example
in the cell P5.
i) MATCH($O5,$A$5:$A$805,0) searches cells from A5 to A805 for the wavelength in O5 and get
a position information (index) of the matched wavelength value. Index function then searches
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CHEM457,
cells C5 to C805 to recall the transmission efficiency at the wavelength (same to the wavelength
in O5) based on the index found in MATCH function.
ii) Check INDEX and MATCH function for your knowledge. The values are normalized by 100 to
calculate efficiency. The dollar sign fixes a low or a column.
iii) Use the same function to calculate transmission efficiency for each filter in column from R to U.
You need to correct the column shift when you paste and copy the function.
iv) The normalized transmission efficiency is platted in column from O to U.
e) Plotting transmission spectra. Select the entire data from O5 to U405. Select an appropriate chart
format. For example, Insert > Scatter with Straight Lines in the Charts Tool. Scientist must know
how to create professional quality graphs. This means that a graph should have clear axis labels, units,
ticks, legends, caption, and other necessary information. Make it better than the example shown
below.
Figure 4. Transmission spectra of optical filters.
f) You will be given excitation and emission spectra of two fluorophores that are widely used in
fluorescence microscopy. We have fluorophore A and B (in the excel template). Based on this
information, we will configure the best set of excitation and emission filters for each fluorophore.
g) To do this, plot excitation and emission spectra of fluorophore A and examine based on the selection
criteria shown below.
i) Selection Criteria
(1) The greater spectral overlap between the excitation filter and excitation spectrum of a
fluorophore, the better performance.
(2) The greater spectral overlap between the emission filter and emission spectrum of a
fluorophore, the better performance.
(3) The transmission spectra of excitation and emission filter must have no overlap.
h) Configure the best filter set for fluorophore A and plot it together with fluorophore spectra. NOTE:
We assume that 1) dichroic mirror could be configured at any wavelength with 100% reflectance of
excitation light and 100% transparency of emission, 2) Other parameters such as excitation source
and quantum yield are wavelength-independent. Actually, this is not true for real experiments, but we
make the assumption to focus on the filter selection.
7) End of group works. Copy the group analysis spreadsheet using a USB driver and complete the
following analysis yourself. The works after this point is your own work. Do not share the results one
another.
8) In the lab report, you will need to answer the following questions.
a) Configure and plot the best filter set for fluorophore B.
b) When you replace the best emission filter that you just determined with filter #1, how many percent
of the effective brightness (relative brightness) would be decreased? To answer this question, you
!20
CHEM457,
need to calculate the effective brightness (ISignal) with the best filter set (the best excitation and
emission filter), and also with the replacement (the best excitation and replacement emission filter,
#1). Use the equation below. You can do numerical summation (Σ) instead of integral to calculate the
relative brightness. The template is provided for calculation (see the section 9 below).
i)
Effective brightness is proportional to a spectral overlap between a fluorophore and filter’s
transmission efficiency, as described the equation (1) below:
!
.
Here, T(λ) is transmission efficiency of excitation or emission filters and F(λ) is excitation or emission
spectra of a fluorophore.
ii) For example, if you estimate an effective brightness for the best filter set to be 50, and 30 when
the best emission filter is replaced with filter #1, the % decrease is (50-30)/50 *100 = 40%.
9) The analysis template and an example are provided for the calculation of the effective brightness (ISig)
i) See the second tab of FM2_Analysis template_Filter transmission spectra.xlsx
ii) The detailed instruction is on the right side of the sheet.
iii) This is just an example calculated with non-optimal filters.
iv) You need to enter the transmission efficiency (T efficiency) of excitation and emission filters
calculated in the first tab into column G and L in the second tab.
v) The transmission efficiency contains negative values due to absorption by glass where a filter
film is coated. So we need to convert all negative values to zero. This is done in column H and M
by IF function.
vi) Calculate TxF (column I) and sum all calculations into J4. This is the first integration part of the
equation (1) for the excitation filter.
(1) NOTE: You need to multiply T and F at the same wavelength.
vii) Do the same calculation for TxF for the emission filter in O4.
viii) The effective brightness ISignal at P4 is multiplication of J4 and O4.
!21
Chem 457 Fluorescence Microscopy Lab 2 Report
CHEM457,
SECTION:_____, GROUP:_____, NAME:______
Title: Configuring a Fluorescence Microscopy Filter Cube
1. Results
• Configure the best filter set for fluorophore A and present data in Figure 1. Provide an
appropriate figure caption. Discuss why you choose these filters based on the selection
criteria in the lab manual.
• Configure the best filter set for fluorophore B and present data in Figure 2.
• If you replace the best emission filter you determined for fluorophore B with the filter #1, how
many percent of the effective (relative) brightness (Isig) would be decreased? Plot the
spectrum of filter #1 in Figure 2. You need to clearly label the best filter sets and the filter #1.
▪ Answer the estimated decrease in %.
▪ For example, if you estimate a relative brightness for the best filter set to
be 50 and filter #1 to be 30, the % decrease is (50-30)/50 *100 = 40%.
2. Discussion
2.1.Justify your choice of fluorescence filters for fluorophore B.
2.2.Suggest an appropriate cutoff wavelength of the dichroic beam splitter that is compatible
with the best filter set for fluorophore A and B.
2.3.Go to fluorescence protein spectra viewer. Configure excitation, emission, and dichroic
mirror using optical filters and mirrors manufactured by Chroma.
– Spectra viewer link: https://www.fpbase.org/spectra/?
s=250,251&showY=0&showX=1&showGrid=0&areaFill=1&logScale=0&scaleEC=0&scaleQ
Y=0&shareTooltip=1&palette=wavelength
– Chroma link: https://www.chroma.com/products/optical-filters
!22
CHEM457,
Darkfield Microscopy: Visualizing Plasmonic
Nanoparticles and Optical Resolution
Darkfield Microscopy
The dark-field microscopy utilizes a special illumination technique to Figure 1. Comparison between darkfield and
enhance the contrast in unstained samples. It works by illuminating
brightfield microscopy
the sample with light that will not be collected by the objective lens
and thus will not form part of the image. In contrast, the light
scattered from the sample enters the objective and form an image.
This produces the classic appearance of a dark, almost black,
background with bright objects on it.
In darkfield microscopy the condenser is designed to form a
hollow cone of light, as opposed to brightfield microscopy that
illuminates the sample with a full cone of light (Figure 1). In
darkfield microscopy, the objective lens sits in the dark hollow of this
cone and light travels around the objective lens, but does not enter the
cone shaped area. The entire field of view appears dark when there is
no sample on the microscope stage. However, when a sample is
placed, it scatters the illuminating light. The scattered light is
collected by an objective lens and forms an image which appears
bright against a dark background. Therefore, the darkfield is a
scattering-based image contrast method.
The objective of this lab is to 1) visualize plasmonic gold nanoparticles using dark-field microscopy, 2)
examine their spectral properties and 3) quantify the point spread function of a diffraction limited system.
Spectral properties of plasmonic nanoparticles
Plasmonic nanoparticles are noble metal nanoparticles such as gold and silver with a diameter below 100 nm.
They interact with specific wavelengths of light through an optical phenomenon called localized surface plasmon
resonance (LSPR). For more information, watch the following videos:
• Tiny treasure: The future of nano-gold https://www.youtube.com/watch?v=QorK2X7GsVU
• Synthesis of gold nanoparticles https://www.youtube.com/watch?v=3lbOiJgdwFA.
The LSPR of noble metal nanoparticles arises when photons
of a certain frequency induce the collective oscillation of
conduction electrons on the nanoparticles’ surface. This causes Figure 2. UV-visible spectroscopy of gold metal
selective photon absorption, efficient scattering, and enhanced nanoparticles
electromagnetic field strength around the nanoparticles. The
color of gold nanoparticles is determined on wavelengths of
light that interact the particles. Figure 2 is a typical UV-Vis
spectrum of spherical gold nanoparticles in solution. The gold
nanoparticles we are using strongly scatter wavelengths
around 525 nm. Since dark-field microscopy forms an image
from scattered lights, spherical gold nanoparticles appear
green in a darkfield microscope. However, their solution color
is red because transmitted lights contain mostly wavelengths
greater than 600 nm.
Resolution in a diffraction limited system
In microscopy, the term ‘resolution’ is used to describe the ability of a microscope to distinguish detail. In
other words, this is the minimum distance at which two distinct points of a specimen can still be seen – either by
the observer or the microscope camera – as separate entities. In this lab, we are going to visualize 65 nm gold
nanoparticles using darkfield microscopy and discuss how the optical resolution is defined. The concept of
resolution is universally applied to all other types of microscopy techniques. Let me first ask this question.
What size would the particle look under the microscope? Would it be about its actual size as we have seen with
other samples? The answer is No. They will appear in a much bigger size—a few hundreds of nanometers. In
principal, 20, 40, 65, and 100 nm particles will appear in the same size. Isn’t it interesting? This means that there
is a fundamental size limit. When light passes through any size aperture (an objective lens in microscopy has a
finite aperture), diffraction occurs. The resulting diffraction pattern, a bright region in the center, together with a
!
23
series of concentric rings of decreasing intensity around it, is
called the Airy Disk (Figure 3). Therefore, in optical
microscopy the optimally focused point of light appears as an
Airy Disc limited by diffraction. The central point of the Airy
Disc contains approximately 84% of the luminous intensity
with the remaining 16% in the diffraction pattern around this
point. This Airy patten is often called the point spread
function (PSF). The central peak reason is approximated to a
two-dimensional (2D) Gaussian function:
f! (x, y) = Ae
−
CHEM457,
Figure 3. Airy Disk, with its central maximum point of
light and the encircling diffractive rings. Left: twodimensional view. Right: three-dimensional view.
2
2
(x − x 0) + (y − y0)
2
2σ
Where !σ is the standard deviation. If one fits the PSF to a 2D Gaussian function, the position of the particle
!(x 0, y0), the max intensity (A), and the standard deviation can be determined. If you need to refresh the Gaussian
function, see Figure 4.
Figure 4. Gaussian distribution. One standard
deviation, or one , plotted above or below the
average value on that normal distribution curve,
would define a region that includes 68 percent of all
the data points. The full width at half maximum
(FWHM) is the width of a line shape at half of its
maximum amplitude.
Figure 5. Superposition of two Gaussian peaks at different separations.
The resolution—the minimum distance at which two distinct points can be distinguished—depends on the
width of the PSF. Suppose you have two PSF (for example, images of two gold nanoparticles). When the gold
nanoparticles are too close to each other, they will appear as a single entity (Figure 5. Top left, when the distance
between two centroid is 2!σ). To be resolved, the separation should be at least 3!σ (top right). This 3!σ is the
experimental (or practical) resolution—it can be determined by a fitting a PSF of a single gold nanoparticle to a
2D Gaussian function.
Theoretical resolution for a given imaging system can be calculated by Abbe’s diffraction formular:
d= λ/2 NA
Where λ is the wavelength of light used to image a specimen. NA is the numerical aperture of an objective. If
using a green light of 514 nm and an oil immersion objective with an NA of 1.45, then the (theoretical) limit of
resolution will be 177 nm. Typically, the practical resolution is poorer than the theoretical one.
Although the fundamental resolution of optical microscopy is about a few hundred nanometers, the position
of individual point emitter (a single gold nanoparticle in this lab) can be determined with much higher precision
—often with several nanometer precision. The position can be determined by fitting a PSF to a 2D Gaussian
function (Figure 6). Localization of each emitter essentially amounts to a summed measurement of its lateral
position, where each photon that can be detected from any specific emitter constitutes a single measurement of
that position. Thus, as more photons are detected, the result will be increasingly better localization precision. The
localization precision can be described by the following equation:
σ
! ∆x y =
N
Where !σ is the standard deviation of the point spread function and N is the number of photons collected.
!24
CHEM457,
Figure 6. Determine the position of a single emitter. Raw data (a) is fitted to a 2D Gaussian function (b). The centroid (position) can be
determined with much higher accuracy compared to the optical resolution.
Experiments
Important notice
4) Wash your hands thoroughly before experiments. Please be careful not to touch any lenses including
objectives, condenser, and eyepieces. You can touch their bodies, but not lenses. If you do so, notify the
instructor of it.
5) Whenever you change objectives and load/unload a sample, lower the sample stage. If you don’t, the
objective may hit the stage and be damaged.
Microscope specification
3) Model: T660C-DKO-IRIS (AmScope), Labeled with DF A or B
4) Can be used in either a bright field microscopy or dark-field microscopy mode
5) Camera: CM3-U3-31S4C-CS, Color camera, 3.2 MP (2048 x1536), 55 FPS, SONY IMX265, FLIR
6) Oil dark-field condenser NA 1.36-1.25
7) NA of objectives: 10x NA 0.25. 40X NA 0.65.
Overview of the experiment
6) Carefully examine the optical and operational components of the microscope
7) Align the condenser
8) Acquire images
9) Lab report preparation.
a) Getting started: examine microscope parts and get started
4) The microscope should already have the darkfield condenser installed by an instructor.
5) Examine labels on the microscope.
6) Turn on the light for brightfield (darkfield) imaging: Switch on the light source.
7) Light intensity control: Adjust the light intensity using the light control knob. Finish at the highest
intensity setting.
8) Sample stage: Move the sample stage in x/y directions by turning the x/y knob of the stage control arm.
9) Focus knobs: Raise and lower the sample stage by turning the coarse and fine focus knobs.
10) Condenser knob: Raise and lower the condenser by turning the condenser knob.
11) Open and close field diaphragm (iris). Fully open the field diaphragm.
12) Turning the nosepiece to change objective lenses. You must lower down the stage before moving the
nosepiece.
13) Push in and pull out the beam splitter that controls the light path – In: eyepiece only; out: eyepiece and
camera. Finish with OUT (camera and eyepiece dual mode).
14) Finish with basic operation practice with the following setting.
– sample stage lowered. Use the focus knob to adjust
– sample stage in the center. Use x/y stage moving arm to adjust
– 10x objective lens in the imaging position
– the field diaphragm fully open
– the beam splitter in the “out” position.
– Light source intensity set to the minimum level.
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b) Immobilize gold nanoparticles on a glass substrate (chemical principles behind the sample preparation)
”
Figure 1. Immobilization of gold nanoparticles on a poly-L-lysine-coated glass substrate.
We will image gold nanoparticles using the darkfield microscopy. To do this, we will immobilize gold
nanoparticles on the surface of a glass substrate via electrostatic interaction. Gold nanoparticles were
synthesized by reducing chloroauric acid (HAuCl4) with sodium citrate. Sodium citrate serves as a reducing
agent and also a stabilizer. The gold nanoparticle surface is negatively charged due to citrate anions adsorbed
on the nanoparticle surface. The negatively charged surface prevents the nanoparticles from aggregation. The
surface of the glass substrate is also negatively charged. Therefore, we have to convert the surface charge of a
glass from negative to positive to facilitate electrostatic interaction between the nanoparticles and glass
substrate. This can be done easily by coating a glass substrate with poly-L-lysine. So, we will first coat the
glass substrate with poly-L-lysine, then add gold nanoparticle solution to adsorb on the glass. The sample will
be prepared in a microfluidic imaging cell as described below.
c) Sample preparation
1) Prepare a flow cell
Note: the example of a flow cell assembly cell is available at the front bench. Talk to Dr. Lee and take a
look at it before you start.
!
Figure 2. Schematic drawing of a flow cell.
i)
Apply double layers of 7-cm-long-double side sticky tapes separated by 5 mm onto 1-mm-thick glass
slide.
ii) Cover 0.17-mm-thick-cover glass.
iii) Press the edge of the cover glass to seal the chamber.
iv) Ready to use
2) Coating a glass with poly-L-lysine (PLL)
i) Introduce 60 uL of 0.1% poly-L-lysine solution into the flow cell as shown in figure 3. The solution
will flow in spontaneously by capillarly action.
!
Figure 3. Adding poly-L-lysine solution into a flow cell.
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ii) Incubate for 5 min.
iii) Wash with DIW three times (Ask Dr. Lee to show this, if this is not clear)
(1) Apply 80 uL of DIW into the one end where you have applied the PLL solution (inlet) while
soaking up the PLL solution from the other end of the chamber (outlet).
(2) This will allow you to introduce DIW to wash sample and remove the existing PLL solution from
the chamber.
Note: Use a tweezer to handle the tissue. The PLL is not classified in hazards. However, wash with
water in case of skin contact.
(3) Repeat the washing two more times.
3) Introduce gold nanoparticles.
i) Prepare one sample for each size of the gold nanoparticle (65 and 80 nm).
ii) Prepare 60 uL of 5x dilution of gold nanoparticle solution in DIW. Total diluted volume should be 60
uL. Calculate the volume of gold nanoparticle solution and DIW required to prepare the diluted
solution.
iii) Get a piece of tissue ready. Apply gold nanoparticle solution in the same way you washed with DIW.
iv) Incubate for 10 mim.
v) Wash the flow cell with 80 uL of DIW three times.
d) Visualize 65 nm gold nanoparticles with a 10x objective Lab Report Figure 1
1) Setting up the microscope and sample loading
i) Set the 10x objective lens.
ii) Login computer
(a) ID: COS-SMBClabcsl222
(b) PW: ChemSMBCcsl222@
iii) Open Point Grey FlyCap2 on desktop (NOTE: Do NOT open Micro-manager). Select the camera
then click OK.
iv) In the menu bar, go to Settings > Toggle Camera Control Dialog.
v) Set the camera setting as follows
(a) Uncheck all check boxes except for Absolute Mode and Power.
(b) Shutter 30 ms (in FlyCap2, shutter is same as exposure time in Micro-manager)
(c) Gain 30
vi) Apply 70 uL of oil on the top lens of the darkfield condenser. Avoid bubbles. If you have bubble, you
need to clean and reapply the oil. You may lower the condenser height to have enough room.
(a) Tips on how to avoid bubble formation: The oil is very viscous. When you pipette up the
oil, slowly release the pipet plunger and wait for 10 s before remove the tip from the oil
solution.
vii) Place the sample on the stage.
viii)Raise the darkfield condenser to the max level using a condenser knob. The oil on the condenser top
lens should now contact the back surface of the slide.
ix) Increase the light intensity to the max level.
x) If your camera is not in live, click the play button !
to start.
xi) Adjust the focus until you see gold nanoparticles.
(a) Note: you may need to adjust the shutter and gain to obtain the optimal image.
2) Align the condenser
i) As long as you fully raise the condenser, the height should be right.
ii) The condenser alignment of the darkfield microscopy is not the same as bright field microscopy. You
do NOT visualize the blade edges of a field diaphragm. Keep the field diaphragm wide open.
iii) Your condenser may not be aligned in x/y. Use the two top silver know to achieve even illumination
thought the image. Do not use the bottom black caped knob.
3) Image acquisition
i) We may need to just Shutter and Gain in Toggle Camera Control Dialog to avoid camera
saturation. The gold nanoparticles should appear green. When Shutter and Gain are too high, the
particles appear in white or yellow color. Reduce either Shutter or Gain until you see green particles
with dark black background.
ii) Capture an image and keep the information about the shutter and gain to prepare your lab report.
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iii) Click the recording button !
to open recording settings.
iv) Click “Browse” in the file name.
v) Select the file path and enter the file name.
vi) Capture 1 frames
vii) Image Format TIFF
viii)Compression Method: None
ix) Click Start Recording to save file.
x) Note: You need image and its acquisition conditions (e.g. shutter and gain value) for your lab report.
Go to your directory and confirm your image.
e) Visualize 65 nm gold nanoparticles with a 40x objective. Lab Report Figure 1
1) Lower the sample stage (Do Not lower the condenser).
2) Change the objective to 40x.
3) Focus the sample.
4) Adjust the Shutter and Gain to obtain the optimal image. Make sure you do not saturate the camera.
5) Capture an image and keep the information about the shutter and gain to prepare your lab report.
Note: You need 40x image and its acquisition conditions (e.g. shutter and gain value) for your lab report. Go
to your directory and confirm your image.
f) Take 10x and 40x images for 80 nm gold nanoparticles. Lab Report Figure 1
Note: You need to find the optimal shutter and gain values for imaging. Keep the shutter and gain values for
preparing the lab report. You will take at least 4 images in total (10x and 40x for each, 2 samples).
g) Finish the experiments
1) Lower the stage.
2) Lower the condenser.
3) Discard the sample.
4) Clean up the oil on the condenser and other parts. Ask to get lens paper if it is not available next to the DF
microscope.
5) Close the software.
Chem 457 Darkfield Microscopy Lab Report
SECTION:_____, GROUP:_____, NAME:______
Title: Visualizing gold nanoparticles and optical resolution
1. Results
• Figure 1. Present 10x and 40x gold nanoparticle images with a 5 micrometer scale bar.
o The calibrated pixel size of 10x objective: 0.85 µm/pixel; 40 x objective: 0.21µm/pixel.
o See this page for how to add scale bar: https://www.swarthmore.edu/NatSci/nkaplin1/
scalebar.htm
▪ Note: As we already know the calibrated pixel dimension, we do not need to do
step 1-3. You can directly set the scale starting with step 4. For example, for 10x
objective the distance in pixel is 1. The known distance is 0.85 µm.
▪ A scale bar example is shown below
•
•
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!
Calculation 1. Calculate the theoretical optical resolution for 10x and 40x objective. NA of
an objective can be found in the lab manual. Use the peak wavelength of the gold
nanoparticles in a UV-Vis absorption spectrum. See the introduction for more information.
Determine the experimental resolution for 65 and 80 nm particle images obtained with a 40x
objective (no need to do 10x). Figure 2. Present a frequency histogram (from minimum
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20-30 particles for each size) with average and standard deviation values. Display 65 and
80 nm in the same graph.
o To determine the experimental resolution, you need to estimate σ
! by fitting point
spread functions (PSF) of individual gold nanoparticles. The fitting can be done
using an ImageJ plugin called GaussFit OnSpot. You can download the plugin on
this page: https://imagej.nih.gov/ij/plugins/gauss-fit-spot/index.html
o The user manual is here: https://imagej.nih.gov/ij/plugins/gauss-fit-spot/
GaussFit_OnSpot.pdf
o The plugin file and manual are also available on Canvas.
o Once installed, you can find GaussFit OnSpout under Plugins in the main menu.
o It should not take more than 15 min to install and learn how to use. If you spend an
unusually large amount of time, contact Dr. Lee.
o As described in the manual, you need to select particles to analyze using the multi-
o
o
o
o
point tool. Hold on the point tool !
then change it to multi-point tool !
. Then
select nicely focused, well isolated single nanoparticles (minimum 20-30
particles). Zooming in image is recommended before selecting particles. Do not
select particles that are too close to each other. For example, do not select
particles less than about 8 pixels of separation with their neighbors.
In GuassFit OnSpot, set Shape to Circle, FitMode to NelDerMead, Ractangle
HalfSize to a bit greater (~1 pixel) than radius of your spots.
Use pixel size described above, and cPCF to 1.0.
Base Level set to 0 photons.
Use the sigma !σ values to calculate the experimental resolution. See the
introduction for how to calculate the experimental resolution from σ
! .
2. Discussion
a. Compare and discuss the experimental resolution of 65 and 80 nm gold nanoparticles imaged
with a 40x objective. Do they appear in the same size or in different size? Why? (Discussion of
figure 2)
b. Depending on the particle size and shape, gold nanoparticles interact with different wavelengths
of light. A is a 65 nm spherical gold nanoparticle that we measured in the lab, and B is a 100 nm
gold nanoparticle with bumpy surface morphology. Note: Optical Density is the same as
Absorbance.
UV-VIS spectra of (A) spherical 65 nm gold nanoparticles and (B) urchin-shaped gold
nanoparticles (“spiky gold”).
What color would 100 nm gold nanoparticles (B) appear in dark field microscopy? Justify the
color of the gold nanoparticle based on the imaging contrast principle of dark field microscopy
and UV-Visible spectrum shown below.
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To prepare the lab report, copy and paste the questions and type your answers. There is no lab report template.
Make sure that you include your name, Section, Group, and Red ID.
Characterization of gold nanoparticle Part I:
Optical Properties of Gold Nanoparticles
Background Information
Nanoparticles have diameters of 1-100 nm, so a 1 nm particle has a diameter of 1 x 10-9 m. Nanoparticle
research is currently a very active field of investigation due to a large variety of applications spanning from
medicine (1), optics and electronics (2) and chemical and biological detection and measurement (3).
Nanoparticles are special because they display characteristics that are different from those of the same material in
bulk form. In fact, one can fine tune physical and
optical
properties of nanoparticles by controlling their size
a n d
shape.
You are familiar with the shiny, metallic
appearance of bulk gold and silver.
However,
reducing the dimensions of the particle drastically
changes the appearance as a result of the way it
interacts with light. Gold nanoparticles with
diameters of 25 nm absorb green light and appear red
i
n
color. Silver nanoparticles absorb violet light and are
yellow.
The ability of nanoparticles to absorb or scatter light
is not
new knowledge. Artisans as far back as the times of
Mesopotamia used nanoparticles to generate a
glittering effect on ceramic pots and artists in
medieval times used them for stained glass. Because
nanoparticles are stable, the red and yellow color in
these
windows remains today (see the figure on the right).
Nanostructures like those that produce bright,
shimmering colors on butterfly wings are composed
o
f
multiple layers with air gaps in between that refract,
diffract
and reflect light generating luminous colors (4).
Nanoparticle shape also affects its optical properties.
F o r
example, spherical gold nanoparticles absorb in the
5 0 0
nm spectral region while irregularly shaped nanorods
a n d
nanostars absorb in the near-infrared (5).
A large area of application of nanoparticles is in
t h e
development of optical sensors for detection and
measurement of a wide array of analytes (3).
Spherical gold nanoparticles change color from red to
b l u e
depending whether they are dispersed or aggregated.
Thus,
any ion, small molecule or even protein that can
trigger
gold nanoparticles to aggregate or disperse can be
detected.
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Figure 1. Example of colorimetric sensing of metal ions using gold nanoparticles functionalized with
chelating agents. Monodispersed gold nanoparticles aggregates via chelating agents (green) and turn its
solution color from pink to blue.
For example, sensors have been developed that rely on gold nanoparticles coated with chelating agents. In
the absence of the target ion, the nanoparticles are in their dispersed state and appear red. In the presence of the
specific ion, the interaction between the chelating agent and the ion bring the nanoparticles together, shifting their
color to blue. Similar approaches using different functional groups on the nanoparticles have extended this
detection approach to small organic molecules, oligonucleotides and proteins (3).
The color change of the gold and silver nanoparticles illustrates an important concept about nanoscale
science. Chemical and physical properties such as color, conductivity, and reactivity do not depend on the
identity of the substance but on the size of the particle.
Synthesis of Gold Nanoparticles
There are several different ways by which gold nanoparticles can be synthesized but the most common
reaction involves the reduction of tetrachloroauric acid (HAuCl4) to gold (Au) using trisodium citrate
(Na3C6H5O7) (6). This is a red-ox reaction where gold in the tetrachloroauric acid is reduced to elemental gold
while the citrate (C6H5O7 3–) is oxidized to dicarboxy acetone (C5H4O5) (7). Figure 3 shows the structures of
these species. This reaction was first described by John Turkevich in 1951 (8) and the mechanisms of particle
formation under different experimental conditions are well discussed in the literature (9).
Q1. What is the oxidation number of gold in HAuCl4?
Q2. Write the half reaction for the reduction of HAuCl4 to Au.
(a)
(b)
(c)
2.Chemical structure of citric acid (a); trisodium citrate (b); dicarboxy acetone (c)
Excess citrate ions not involved in the red-ox reaction are also adsorbed on the surface of the particles, thus
playing a role in stabilizing the nanoparticles. They are called “capping” agents. Adsorption of citrate ions gives
the gold particles an overall negative charge. Mutual repulsion of the small, negatively charged particles keeps
them suspended in the solution and prevents them from coagulating to form larger particles that might eventually
settle out of solution. This suspension of the gold nanoparticles is known as a colloid.
Since citric acid is a triprotic acid with distinct pKa values of 3.2, 4.8 and 6.4, pH will affect the chemical
equilibrium involving the dissociation of the three hydrogens (10).
Q3. Above what pH value will citric acid (C6H8O7) be completely dissociated to citrate (C6H5O73–)?
From an application standpoint, it is very important that particle size distribution be uniform. For example, if one
was to develop a colorimetric sensor for a given analyte based on the color shift of gold nanoparticles, the optical
characteristics of the nanoparticles would have to be highly reproducible to ensure repeatability of the analytical
method. As shown in Table 1, Size strongly affects the particle wavelength of maximum absorption as well as the
molar extinction coefficient which, in turns, defines the sensitivity of the analytical method.
Thus it is important to understand how different synthetic conditions influence the size and uniformity of size
distribution. Controlling the size and size distribution is necessary to control the optical properties of the
nanoparticles.
Exploring fundamental relationships in spectrophotometry
Solutions of gold nanoparticles appear colored which means they are absorbing light in the visible range.
Since you will be investigating the optical properties of gold nanoparticles using a technique called
spectrophotometry, we will briefly review how a spectrophotometric measurement takes place.
When a beam of electromagnetic radiation passes through a sample, most of the radiation is transmitted but,
at specific wavelengths, the radiation may be absorbed by chemical constituents within the sample. For an atomic
or molecular species, the absorption of light causes valence electrons to be excited from lower to higher energy
!
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CHEM457,
states. For this transition to occur, the energy provided by the radiation has to match the difference in energy
(DE) between the lower and higher energy states.
In the gold nanoparticle lab part 2, we will quantify the concentration of gold nanoparticles from a UV-Vis
spectrum. In this lab, we will build a fundamental knowledge about determining concentration using
spectrophotometry.
The amount of light transmitted through a sample is measured by the transmittance and is represented by
the equation:
T = P/P0.
where P0 represents the radiant power of the beam before the sample and P the power after the sample.
Another commonly used measurement is the absorbance which is related to the transmittance through a
logarithmic relationship:
A = – log T = – log P/P0 = log P0/P
Relationship between absorbance and concentration
It is possible to make quantitative measurements using UV-VIS spectrophotometry because of the linear
relationship between absorbance and concentration. This relationship is known as Beer’s law and is represented
by the equation:
A = ebc
where
A is the absorbance
e is the molar extinction coefficient (with units of M-1cm-1)
b is the path length of the cuvette (cm)
c is the concentration (Molarity)
The changes in absorbance, A, with changes in concentration of a sample are measured at lmax which is the
wavelength of maximum absorption in the spectrum of the compound being analyzed. By plotting A as a function
of varying concentrations of the analyte, a regression curve can be established. A representative plot of A versus c
is shown below.
”
How could we use this plot to determine the molar extinction coefficient of the analyte being investigated?
Q4. The following data shows the absorbance of molecule X at different concentrations. Determine the
extinction coefficient. The path length of the Cuvette is 1 cm. Describe how you determined the extinction
coefficient in detail.
Concentration [M]
Absorbance
1.00E-05
0.6
2.00E-05
1.8
4.00E-05
2.8
6.00E-05
4.9
8.00E-05
6.0
The molar extinction coefficient can be linked to the intensity of light absorption. For example, transition
metals and dyes appear brightly colored because some transitions in the visible range are highly probable and
have very large extinction coefficients. For example, the extinction coefficient of the bright Bromothymol Blue at
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615 nm is approximately 3.5 x 104 M-1cm-1. Gold nanoparticles are also intensely colored and their color as well
as its intensity is controlled by their size (Figure 2).
!
Figure 3. Colors of monodispersed metal nanoparticles in various shapes and sizes
Table 1. Effect of particle size on maximum wavelength of absorbance and molar extinction coefficient of gold
nanoparticles.
Diameter (nm)
Peak Wavelength (nm)
Molar Extinction Coefficient (M-1cm-1)
5
515-520
1.10 x 107
10
515-520
1.01 x 108
15
520
3.67×108
20
524
9.21 x 108
30
526
3.36 x 109
40
530
8.42 x 109
50
535
1.72 x 1010
60
540
3.07 x 1010
80
553
7.70 x 1010
100
572
1.57 x 1011
S o u r c e : h t t p : / / w w w. s i g m a a l d r i c h . c o m / m a t e r i a l s – s c i e n c e / n a n o m a t e r i a l s / g o l d nanoparticles.html#sthash.kwu2f19v.dpuf
Q5. What trend can be identified in the molar extinction coefficient value as particle size increases?
Q6. What color would a nanoparticle solution that absorbs in the blue-green region of the electromagnetic
spectrum appear?
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!
Q7. If you know the extinction coefficient, e, of the nanoparticle, The UV-Vis spectrum shown above can be
used to determine the concentration of the nanoparticle using Beer’s law represented by the equation: A =
ebc. At which wavelength should the absorbance be measured? You can answer the approximate
wavelength. Provide justification.
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Characterization of Gold Nanoparticles Part II:
Determining concentration of nanoparticles
using Transmission electron microscopy and UVVis spectroscopy
Determination of the size of nanoparticles from
Purpose: The purpose of this exercise is to determine the average diameter of gold nanoparticles using a
Transmission Electron Microscope (TEM). In addition, the size distribution will be evaluated. Then we will
determine the concentration of gold nanoparticles from a UV-Vis absorption spectrum.
Learning Outcomes:
At the end of this assignment you will be able to
1. Analyze TEM images to determine the average particle diameter. You will extend the ImageJ-based
quantitative image analysis skills beyond optical microscopy.
2. Determine the size distribution for each nanoparticle preparation previously synthesized.
3. Determine the concentration of nanoparticle from a UV-Vis absorption spectrum.
Figure 1. HRTEM images of gold nanoparticles synthesized by reducing tetrachloroauric (III) acid by
sodium citrate.
The images shown in figure 1 were collected using a High Resolution Transmission Electron Microscope
(HRTEM) on nanoparticles prepared by reducing tetrachloroauric (III) acid by sodium citrate. To quantitatively
analyze the images and extract information about particle size and size distribution, you will use a free software
called ImageJ (FIJI). The first task will be to determine the Feret’s diameter of the particles. See the appendix
of this document (at the end) for step-by-step procedure to determine the particle diameter. Go the Canvas module
page, Chracterization of Gold Nanoparticle PART 1 and 2. Download a HRTEM image of gold nanoparticles.
Next, load the image in ImageJ and determine the Feret’s diameter.
Q1. What is the average value of the Feret’s diameter you obtained from the analysis of the image?
NOTE: The Appendix at the end of this document provides step-by-step procedure to analyze the diameter of
particles in a TEM image.
Next, you will explore the particle size distribution. The output from the ImageJ analysis will list each detected
particle and its diameter. Count all the particles of a given diameter (you may want to group them in brackets
within 0.5 or 1 nm) and, using a spreadsheet, compile a chart reporting the probability of occurrence of each
particle of a given diameter. This type of graph is called histogram.
Q2. What is the standard deviation?
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Q3. Display a probability distribution of the Feret’s diameter of gold nanoparticles.
Estimating the concentration of nanoparticles from the particle size data
Purpose: The purpose of this exercise is to estimate the molar concentration of nanoparticles based on the particle
diameter measured by TEM and a UV-Vis absorption spectrum.
Learning Outcomes:
At the end of this assignment you will be able to
1. Calculate the molar concentration of any nanoparticle preparation using the Feret’s diameter estimated
through TEM image analysis and a UV-Vis absorption spectrum.
2. Use a diameter determined by TEM image analysis and calculate the amount of gold atoms per
nanoparticle.
Assignment:
The gold nanoparticle shows different interactions with light depending on their size and shape.
As shown in the figure 2, the absorption maximum (lmax) increases from 520 nm to 570 nm for 20 nm and 100
nm spherical gold nanoparticles, respectively.
!
Figure 2. Gold nanoparticle size-dependent surface plasmon resonance. Note the red-shift of the absorption
maximum as the gold nanoparticle size increases.
The table 1 shows the extinction coefficients of gold nanoparticles with different sizes. The concentration of gold
nanoparticles can be determined from the absorbance value using the Beer-Lambert law, as we discussed in the
lab PART 1.
Table 1. Effect of particle size on maximum wavelength of absorbance and molar extinction coefficient of gold
nanoparticles.
!36
Diameter (nm)
lmax (nm)
Molar Extinction Coefficient (M-1cm-1)
5
515-520
1.10 x 107
10
515-520
1.01 x 108
15
520
3.67×108
20
524
9.21 x 108
30
526
3.36 x 109
40
530
8.42 x 109
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50
535
1.72 x 1010
60
540
3.07 x 1010
80
553
7.70 x 1010
100
572
1.57 x 1011
Gold nanoparticles are synthesized and suspended in solution. You can find a UV-Vis spectrum of the gold
nanoparticles for which you just determined a diameter. Download a UV-Vis spectrum from Canvas, and
determine the concentration of gold nanoparticles.
Q4. What is the concentration of gold nanoparticles? Present equations.
Note: you may select the extinction coefficient for the gold nanoparticle diameter that is closest to the diameter
you determined. The path length of a UV-Vis cell was 1 cm.
Q5. What is the number of gold nanoparticles in 10 uL of solution? Present equations.
The average number of gold atoms per nanoparticle can be calculated if the diameter of a nanoparticle is known.
In the specific case of gold, Liu and coworkers (11) determined the following relationship between the average
number of gold atoms (N) per nanoparticle and the particle diameter (D):
N= !
π (19.3
cm 3 )
g
D3
6(197 mol )
g
(eq. 1)
This equation assumes a spherical shape and a uniform face-centered cubic (fcc) structure. In equation 1, 19.3 g/
cm3 is the density for fcc gold and 197 g/mol is the gold atomic mass.
Q6. How many moles of gold atoms are present per gold nanoparticle? Present equations.
Q7. What is the number of gold atoms per gold nanoparticle? Present equations.
APENDIX. Image J Processing: Determining Particle Diameter using ImageJ (FIJI)
The following procedure outlines a standard approach to using Image J with TEM images. It is recommended
that images be saved in .TIFF format.
Additional information, including examples and tutorials, can be found on the ImageJ homepage (https://
imagej.nih.gov/ij/) in the Documentation link.
1. Double click on the (FIJI or ImageJ) shortcut on the desktop to open the image processing software.
2. Load the TEM image. Do this by selecting “File” and then “Open” or drag-drop the file onto the main
window of FIJI.
3. The software from modern TEM instruments will embed a scale bar on the image in order to give a
reference to the size of objects in the image. We will use this to tell the software how big each pixel in
the digital image is, which the software can then use to calculate sizes in a more automated manner. This
is the same as what we did to measure the length the spike protein of SARS-CoV-2.
4. Use the zoom tool (looks like a magnifying glass) or the + and – keys on the keyboard to enlarge the
image until the scale bar in the image almost fills the screen. Pan around the image using the tool that
looks like a hand.
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!
5. Select the line tool from the tool bar at the top of the screen and draw a line over the scale bar (the line
should extend exactly the same length as the scale bar).
!
6. Select “Analyze” and then “Set Scale”.
!
7. Remove any prior measurements by selecting “Click to remove scale”.
8. In the box labelled “Known distance”, enter in the value on the scale bar in your image. The distance in
pixels should already be filled from the line drawn. In the units box, enter the appropriate units, in this
case nanometers or nm. Leave the pixel aspect ratio as the default value of 1 (this just means that the
horizontal and vertical sizes of the pixels are the same). Click “OK”.
9. Next be sure to zoom back out until you can see the entire image (hold control while clicking with the
magnifying glass to zoom out or right click), and then select the region of interest (where you have
nanoparticles, but excluding the margins and scale bar). Do this by clicking on the rectangle tool and
surrounding the area you want to use. You should be able to define a yellow rectangle region.
!
10. Select “Image” and then “Crop”.
11. You next need to adjust the image so that the software can readily distinguish between a nanoparticle and
the background. To do this, you will first adjust the brightness/contrast of the image. Select “Image”
followed by “Adjust” and then “Brightness/Contrast”. Sometimes the “Auto” feature is sufficient, but if
not you can also play around with the sliding bars. Try to adjust them so that the nanoparticles really
stand out and the background is mainly white, but make sure the size of the dots doesn’t change.
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!
12. Select “Image” followed by “Adjust” and then “Threshold”. Make sure you select Red (B&W should
work, but Red is easier to visualize). Move the sliders until the particles you wish to analyze are red, but
none of the background is red. Make sure the size of the dots doesn’t change. This will set the background
to be completely white to create a firm visual boundary between the nanoparticle and the background.
13. Click “Apply”.
!
14. Select “Process”, followed by “Binary” and then “Make Binary”.
!
15. We are about to measure the size of the particles, but first, we need to tell the software what
measurements we want. Go to “Analyze” followed by “Set Measurements”. Check the box “Ferret
Diameter”, but none of the others. Click OK.
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!
16. Finally, we are ready to have the software measure the size of all the nanoparticles. To do this, select
“Analyze” followed by “Analyze Particles”.
17. Check the boxes “Display Results” and “Exclude on Edges” and “Summarize”. The default size from 0
to infinity is a good starting point, although it can be beneficial to put an upper and lower limit to prevent
errant inclusions. For circularity, a range of 0.6 to 1 is typically acceptable.
!
18. Select OK, and two tables with your results will appear, a summary and complete table of each particle.
19. Look through the results table and watch for unrealistic particle sizes (for example if the area is on the
order of 1×10-4 nm2, this makes no physical sense, it results from background noise being counted as a
particle). You can remove these errant points by right clicking on them and selecting “clear”.
20. For each window, select “File” followed by “save as” to save the table as a spreadsheet. End your
filename with “.txt”.
21. While the complete data table is nice, it is challenging to visualize the results in table form. Instead, you
can create a histogram which shows the size distribution of the particles. In the “Results” data file, the
column labeled with “Feret” shows the Feret diameter for each particle; the Feret diameter robustly
represents a diameter of spherical objects. So, in our case, we can use the Feret diameter as a diameter of
gold nanoparticles.
!40
CHEM457,
The diameter of an object measured with a caliper is sometimes called the caliper diameter; it is
the same as Feret diameter.
22. Plot a probability distribution (histogram) of the diameter (Feret diameter) of gold nanoparticles.
!41
CHEM457,
Appendix of FM1: Single Particle Analysis using
ImageJ (Fiji)
This manual aims to provide step-by-step analysis processes to calculate diffusion coefficients and brightness of
fluorescence microsphears using Fiji. You have to use TrackMate to create data for further analysis in Excel.
Important notes
a. READ CAREFULLY. Don’t skip.
b. For the practice analysis,
a. Enter the exactly same parameters presented in the black text region.
b. You should get the exactly same results shown in the video.
c. For the fluorescence microscopy #1 lab analysis (the actual data you acquired),
a. The values in BOLD BLUE could be different for your data. Change accordingly.
b. Read CRITICAL with extra attention. It includes crucial guides on how to set the parameters.
d. You should have some basic backgrounds of statistics such as histogram and probability distribution—not
drawing skill. Those have been taught in many other classes such as CHEM410A/B. Read this webpage to
refresh yourself: https://statistics.laerd.com/statistical-guides/understanding-histograms.php.
e. If you have questions, please send me an email as soon as possible, so I can help you accordingly.
I. Single Particle Tracking using TrackMate
1. Software installation
a. Go to https://fiji.sc/#download and install Fiji.
2. Preimage treatment
a. Open an image file in Fiji.
• If you have a single video file as an image stack, you can simply drag and drop the file onto the Fiji
window.
• (Optional) If you have separate images of the movie (frame-by-frame) you need to drag and drop the
folder that contains the images. This happens if you select “save as separate images” in multidimensional acquisition when you acquire data.
b. Adjust contrast. If you have difficulty in seeing the image because of brightness/contrast, you can adjust it.
Go to Image > Adjust > Brightness/Contrast. End the adjustment by closing the Brightness/Contrast
window. Do NOT Apply.
c. (Optional) If you acquired multiple movies of the same sample because particle density was too low. you
can combine those movies into a single movie and analyze them all together. Open all movies that you
want to combine. Go to Image > Tools > Concatenate. Make sure that all movies are listed in the
Concatenator window. Click Ok to combine. IMPORTANT: CONCATENATE THE SAME
SAMPLES! DO NOT MIX DATA FROM DIFFERENT SAMPLES.
d. Examine the entire frames and remove error frames if necessary.
• Play a movie by clicking play icon located at the bottom left corner of the image window. You can
change the play speed by right-clicking the play icon. Typically, a rate at 20 frames per second is good
to quickly examine the entire image frames. If your movie does not have error images, advance to
step “e”.
• Carefully check if you have error frames. For the demo image, I found all frames after 73rd frame has
stripes. You may have an error image in your first frame. I found some students …