Dr. Anthony S. Stender wishes to acknowledge technical support through the Nanoscale and Quantum Phenomena Institute (NQPI) at Ohio University. This article&#160;was made possible through start-up funding provided to Dr. Stender by Ohio University.

1. Sample preparation with standard glass microscopy slides
<ol>
	<li>Prepare glass microscope slides for sample deposition. 
	NOTE: In some circumstances, it may be more appropriate to store the glass in ultrapure water instead of ethanol. However, storing in water or air makes the glass hydrophobic over time.
	<ol>
		<li>For best results, purchase glass or quartz microscope slides and cover glass.</li>
		<li>Using a scribing pen, place a shallow and short scratch mark onto the center of each glass cover slip.</li>
		<li>Clean all microscope glass, even if it is purchased &#8220;pre-cleaned&#8221;, to remove glass shards, dust, powder, organic residue, and any other contaminants that affect imaging quality or sample deposition. 
		NOTE: This cleaning method below works well for the types of samples described here and avoids the use of harsh chemicals. Harsher chemicals can etch the glass and require more care in handling and disposal.
		<ol>
			<li>Place microscope glass onto storage racks and then into a beaker, or into a staining jar. Do not place microscope glass at the bottom of beakers and other lab glassware without racking, because each piece and surface of microscope glass should be fully exposed to the cleaning agents.</li>
			<li>Pour ~1 mL of liquid detergent (Table of Materials) into the container and top off the container with water. Sonicate for 30 min. 
			NOTE: Once the cleaning process begins, only handle the glass while wearing gloves, to avoid leaving fingerprint residue on the glass.</li>
			<li>Pour out the liquid contents of the cleaning container into a sink. Rinse container several times with ultrapure water to remove all appearance of detergent. Refill the container with ultrapure water. Sonicate the container with microscope glass for another 30 min.</li>
			<li>Repeat the previous step at least once more. Perform additional rounds of sonication in water until it is obvious that all traces of the detergent have been removed.</li>
			<li>Pour out the contents of the cleaning container. Rinse the container with ultrapure water. Refill the container with ethanol. Sonicate microscope glass for 30 min.</li>
			<li>Pour out the contents of the cleaning container into a waste container. Refill with ethanol. Cover the container to prevent loss of ethanol through evaporation. Store the microscope glass in this container until time of experiment. Slides remain clean and usable as long as they remain submerged in ethanol inside a covered container.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparation of nanoparticle solution
	<ol>
		<li>Using a micropipette, remove a 100 &#181;L aliquot of 0.05 mg/mL gold nanoparticle solution from its original storage container and eject the solution into a 1.5 mL centrifuge tube.</li>
		<li>Centrifuge the sample for 10 min at 6,000 x g.</li>
		<li>Remove the supernatant with a micropipette, in order to remove excess surfactant.</li>
		<li>Using a micropipette, place 100 &#181;L of ultrapure water into the centrifuge tube. 
		NOTE: If not all of the supernatant can be removed on the first attempt, repeat the centrifugation and resuspension steps.</li>
		<li>Briefly vortex the sample to resuspend the pellet. Sonicate immediately afterwards for 20 min to fully resuspend and break up nanoparticle aggregates. 
		NOTE: If the sample is not used immediately, it should be sonicated again for 20 min before depositing solution onto microscope glass.</li>
	</ol>
	</li>
	<li>Sample deposition
	<ol>
		<li>Remove cleaned cover slips and microscope slides from their storage containers. Blow dry the glass with pressurized nitrogen or argon.</li>
		<li>Using a micropipette, drop-cast 6 &#181;L of nanoparticle solution from step 1.2.5 onto the cover slip. To spread out the droplet evenly, carefully place a second, larger piece of microscope glass on top of the cover slip, such as a second cover slip or a microscope slide. Avoid getting air bubbles trapped between the two pieces of glass.
		<ol>
			<li>Turn the sample substrate over, and seal off the edges of the cover slip with a narrow line of nail polish in order to prevent evaporation of the medium solution.</li>
			<li>Alternatively, to image the sample &#8220;dry&#8221;, allow the solution to stand for 5&#8211;15 min on the cover slip, before removing the unwanted piece of glass. Gently blow the cover slip dry with pressurized nitrogen or argon.</li>
		</ol>
		</li>
		<li>If possible, image samples immediately after preparation. If that is not possible, store samples in a covered container, such as a Petri dish until imaging.</li>
	</ol>
	</li>
</ol>
2. DIC imaging
<ol>
	<li>Align objective and condenser.
	<ol>
		<li>After placing the sample onto the microscope, find the focal plane with the sample on it. First locate and focus on the scratch mark created earlier. Then fine tune the focus until nanoparticles come into view.</li>
		<li>To determine the accurate placement of the condenser, utilize the Kohler Illumination method.5 Kohler Illumination at high magnification (80x, 100x) is more easily achieved by first setting the Kohler Illumination at a lower magnification, such as 20x. 
		NOTE: Normally, Kohler Illumination does not need to be re-adjusted during imaging of a single sample. However, it is good practice to verify that Kohler Illumination is properly set when switching to a new microscope slide.</li>
	</ol>
	</li>
	<li>Optimize contrast settings.
	<ol>
		<li>Select a region of interest within the sample for imaging. Center the region in the camera&#8217;s field of view and adjust focus as necessary.
		<ol>
			<li>If the microscope has the de Senarmont design, start with the polarizer set near to maximum background extinction and gradually rotate the polarizer towards decreasing background extinction. The background intensity will gradually increase.</li>
			<li>If the microscope does not have a de Senarmont design, start with the optical train set at maximum background extinction. In this case, gradually adjust the objective prism position towards decreasing background extinction. 
			NOTE: The ideal setting is achieved when the nanoparticles reach their greatest intensity difference (i.e., contrast) from the averaged local background value. For plasmonic nanoparticles, optimal contrast is normally achieved with a relatively dark background, thus at settings near maximum background extinction.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Image the sample.
	<ol>
		<li>Turn off room lighting to prevent stray illumination from interacting with process.</li>
		<li>While viewing the nanoparticles with a scientific imaging camera, determine the optimal background level. Using a 10 nm full width at half maximum (FWHM) bandpass filter with its central wavelength co-located with the main LSPR wavelength, view the region of interest. Adjust the lamp intensity or exposure time until the background level is in the range of 15%&#8211;40% of the camera&#8217;s maximum capacity level and no objects within the region of interest exhibit signal intensities that exceed 90% of the camera&#8217;s maximum intensity level. 
		NOTE: The goal of step 2.3.2 is to prevent saturating the sensor when switching between filters. The ideal background level will vary between samples and cameras. Once this step is completed, exposure time can be adjusted but not the lamp intensity.</li>
		<li>Image the sample with a series of bandpass filters that each has a FWHM of 10 nm and that as a whole enable imaging across the entire wavelength range of interest. Ensure that the background intensity remains consistent from image to image (within ~5% of one another) by adjusting the exposure time. After switching filters, re-focus the sample before image capture.</li>
		<li>Save the Images as uncompressed TIFF files and/or in the software&#8217;s native file format, in order to preserve all information.</li>
	</ol>
	</li>
	<li>Rotate the sample.
	<ol>
		<li>After collecting images of the sample at its original position, the sample can now be rotated and imaged at additional orientations in the light path. Perform rotation at regular intervals (e.g., 10&#176; or 15&#176;) across either a 180&#176; or 360&#176; range. 
		NOTE: Rotation requires a rotatable sample stage.</li>
		<li>As in sections 2.1-2.3, adjust camera settings to provide a consistent background level from image to image. 
		NOTE: No adjustment to Kohler Illumination should be made.</li>
	</ol>
	</li>
</ol>
3. Data analysis using ImageJ
NOTE: The following calculations can be performed in a variety of software packages, and sometimes in the native program used to collect the images. ImageJ is a freely available software from the National Institutes of Health (NIH).
<ol>
	<li>Calculate particle contrast or intensity.
	<ol>
		<li>Open the image with ImageJ.</li>
		<li>Select the Rectangle tool and draw a rectangle around the main region of interest.</li>
		<li>On the Tool bar, select Image, then Zoom, then To Selection. The imaging window will zoom in on the selected area.</li>
		<li>On the Tool bar, select Image, then Adjust, then Brightness/Contrast. A new window appears. To enable better viewing of the sample region, adjust the four settings: Minimum, Maximum, Brightness, and Contrast. These adjustments do not alter the scientific data, they merely enable better visibility of the sample region. 
		NOTE: Steps 3.1.3 and 3.1.4 may be performed multiple times and in reverse order.</li>
		<li>Using the rectangle tool again, draw a box around the first nanoparticle to be measured. The box should be only slightly larger than the nanoparticle&#8217;s airy disc.</li>
		<li>On the Tool bar, select Analyze, then Measure. A new window appears that reports the Minimum, Maximum, and Mean Intensities for the pixels located inside of the selected box.</li>
		<li>Drag the box used to measure the nanoparticle to an area immediately adjacent to the particle, where the background contrast is relatively even and no particles or contaminants are present. Retain the original size of the box.</li>
		<li>Use the Measure tool to determine the Mean Intensity for the background area.</li>
		<li>Measure the remaining particles and an adjacent background area for each.</li>
		<li>Repeat the process for each particle in all of the images in the series.</li>
		<li>Export the data to a spreadsheet to calculate the contrast or intensity of each particle, across all wavelengths and angles.</li>
		<li>Calculate each particle&#8217;s contrast, using the following equation13,14,15: 
		<img alt="Equation" src="/files/ftp_upload/59411/59411eq1.jpg" /> 
		NOTE: Using this equation, particle contrast should always be &#62; 0.</li>
		<li>Calculate the particle&#8217;s background-adjusted maximum value by dividing the measured maximum particle intensity by the background mean: 
		<img alt="Equation" src="/files/ftp_upload/59411/59411eq2.jpg" /></li>
		<li>Likewise, calculate the background-adjusted minimum value by dividing the measured minimum particle intensity by the background mean: 
		<img alt="Equation" src="/files/ftp_upload/59411/59411eq3.jpg" /> 
		NOTE: As calculated, the maximum should have a value greater than one, while the minimum will be less than one. It is acceptable to subtract each value by &#8220;1&#8221;, so that the average background is essentially zero, the maximum is represented as a positive value, and the minimum value is assigned a negative value16. This latter approach allows the analyst to separately consider what is occurring along each of the polarization fields, which is useful when studying anisotropic particles.</li>
		<li>To graph the spectral profile at a given nanoparticle position, plot data with the wavelength along the x-axis and the contrast or intensity along the y-axis.</li>
		<li>To graph the rotational profile at a given wavelength, plot the rotation angle along the x-axis and the contrast or intensity along the y-axis.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Pluta, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch+7:+Differential+Interference+Contrast+in.">Ch 7: Differential Interference Contrast in.</a> Advanced Light Microscopy. 2, 146-197 (1989).</li><li>Stender, A. S., Wang, G., Sun, W., Fang, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+Gold+Nanorod+Geometry+on+Optical+Response.">Influence of Gold Nanorod Geometry on Optical Response.</a> ACS Nano. 4 (12), 7667-7675 (2010).</li><li>Stender, A. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+Cell+Optical+Imaging+and+Spectroscopy.">Single Cell Optical Imaging and Spectroscopy.</a> Chemical Reviews. 113 (4), 2469-2527 (2013).</li><li>Mehta, S. B., Sheppard, C. J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Partially+coherent+image+formation+in+differential+interference+contrast+(DIC)+microscope.">Partially coherent image formation in differential interference contrast (DIC) microscope.</a> Optics Express. 16 (24), 19462-19479 (2008).</li><li>Murphy, D. B., Davidson, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch+1:+Fundamentals+of+Light+Microscopy.">Ch 1: Fundamentals of Light Microscopy.</a> Fundamentals of Light Microscopy and Electronic Imaging, Second edition. , 1-20 (2012).</li><li>Stender, A. S., Augspurger, A. E., Wang, G., Fang, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+Polarization+Setting+on+Gold+Nanorod+Signal+at+Nonplasmonic+Wavelengths+Under+Differential+Interference+Contrast+Microscopy.">Influence of Polarization Setting on Gold Nanorod Signal at Nonplasmonic Wavelengths Under Differential Interference Contrast Microscopy.</a> Analytical Chemistry. 84 (12), 5210-5215 (2012).</li><li>Wang, G., Sun, W., Luo, Y., Fang, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resolving+Rotational+Motions+of+Nano-objects+in+Engineered+Environments+and+Live+Cells+with+Gold+Nanorods+and+Differential+Interference+Contrast+Microscopy.">Resolving Rotational Motions of Nano-objects in Engineered Environments and Live Cells with Gold Nanorods and Differential Interference Contrast Microscopy.</a> Journal of the American Chemical Society. 132 (46), 16417-16422 (2010).</li><li>Kelly, K. L., Coronado, E., Zhao, L. L., Schatz, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Optical+Properties+of+Metal+Nanoparticles:+The+Influence+of+Size,+Shape,+and+Dielectric+Environment.">The Optical Properties of Metal Nanoparticles: The Influence of Size, Shape, and Dielectric Environment.</a> The Journal of Physical Chemistry B. 107 (3), 668-677 (2003).</li><li>Mulvaney, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Not+All+That's+Gold+Does+Glitter.">Not All That's Gold Does Glitter.</a> MRS Bulletin. 26 (12), 1009-1014 (2012).</li><li>Maier, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Plasmonics: Fundamentals and Applications. , (2007).</li><li>Sun, W., Wang, G., Fang, N., Yeung, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wavelength-Dependent+Differential+Interference+Contrast+Microscopy:+Selectively+Imaging+Nanoparticle+Probes+in+Live+Cells.">Wavelength-Dependent Differential Interference Contrast Microscopy: Selectively Imaging Nanoparticle Probes in Live Cells.</a> Analytical Chemistry. 81 (22), 9203-9208 (2009).</li><li>Cras, J. J., Rowe-Taitt, C. A., Nivens, D. A., Ligler, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+chemical+cleaning+methods+of+glass+in+preparation+for+silanization.">Comparison of chemical cleaning methods of glass in preparation for silanization.</a> Biosensors and Bioelectronics. 14 (8), 683-688 (1999).</li><li>Augspurger, A. E., Sun, X., Trewyn, B. G., Fang, N., Stender, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Monitoring+the+Stimulated+Uncapping+Process+of+Gold-Capped+Mesoporous+Silica+Nanoparticles.">Monitoring the Stimulated Uncapping Process of Gold-Capped Mesoporous Silica Nanoparticles.</a> Analytical Chemistry. 90 (5), 3183-3188 (2018).</li><li>Murphy, D. B., Davidson, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch+2:+Light+and+Color.">Ch 2: Light and Color.</a> Fundamentals of Light Microscopy and Electronic Imaging, Second Edition. , 21-33 (2012).</li><li>Wayne, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ch+3:+The+Dependence+of+Image+Formation+on+the+Nature+of+Light.">Ch 3: The Dependence of Image Formation on the Nature of Light.</a> Light and Video Microscopy (Second Edition). , 43-78 (2014).</li><li>Stender, A. S., Wei, X., Augspurger, A. E., Fang, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasmonic+Behavior+of+Single+Gold+Dumbbells+and+Simple+Dumbbell+Geometries.">Plasmonic Behavior of Single Gold Dumbbells and Simple Dumbbell Geometries.</a> The Journal of Physical Chemistry C. 117 (31), 16195-16202 (2013).</li><li>Hu, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dark-field+microscopy+studies+of+single+metal+nanoparticles:+understanding+the+factors+that+influence+the+linewidth+of+the+localized+surface+plasmon+resonance.">Dark-field microscopy studies of single metal nanoparticles: understanding the factors that influence the linewidth of the localized surface plasmon resonance.</a> Journal of Materials Chemistry. 18 (17), 1949-1960 (2008).</li><li>Choo, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wavelength-Dependent+Differential+Interference+Contrast+Inversion+of+Anisotropic+Gold+Nanoparticles.">Wavelength-Dependent Differential Interference Contrast Inversion of Anisotropic Gold Nanoparticles.</a> The Journal of Physical Chemistry C. 122 (47), 27024-27031 (2018).</li><li>Funston, A. M., Novo, C., Davis, T. J., Mulvaney, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasmon+Coupling+of+Gold+Nanorods+at+Short+Distances+and+in+Different+Geometries.">Plasmon Coupling of Gold Nanorods at Short Distances and in Different Geometries.</a> Nano Letters. 9 (4), 1651-1658 (2009).</li></ol>

The author has nothing to disclose.

When imaging with DIC microscopy, it is critical to optimize the optical components before collecting data. Even minor adjustments to the polarizer in the middle of an experiment can result in significant impacts to the final data6. Moreover, different materials require different polarizer settings. Although large step sizes were utilized here to demonstrate the effect of polarization angle, in an actual experiment, it is imperative to optimize the polarizer setting within 1&#176;&#8211;2&#176; of the optimal contrast setting. The polarizer setting should also be recorded for future reference. It is also recommended to always work on the same side of the crossed polarizer (0&#176;) point. Switching back and forth does not provide any advantages, but it can lead to confusion, due to a reversal in the shadow pattern.
Next in importance, it is critical to monitor the background intensity when planning to perform spectroscopy. This is best accomplished by adjusting the camera exposure time, or by adding neutral density filters to the light path. Adjusting apertures or lamp intensities can impact the Kohler Illumination and alter contrast values. The background needs to be relatively even across the sample, so that the selection of a background region does not alter the contrast calculation. Sample specimens that are not adjacent to a clean background space should be avoided. Moreover, the background intensity cannot be initially set too high or too low. If the background intensity is set too high, there is an increased risk that some signals will exceed the maximum range of the camera, making it impossible to calculate the contrast in those regions. If the background intensity is set too low, it will be extremely difficult to achieve good contrast between the dark component of the DIC signal and the background signal. Understanding the typical or expected behavior of a sample can aid in selecting the proper background intensity.
Finding the proper focal plane is also essential. One of Nomarski DIC&#8217;s advantages is that it has a shallow depth of field. However, this makes it more challenging to focus on thin samples, such as nanoparticles. With thicker samples, the challenge is in finding the actual focal plane of greatest interest. Many focal planes may be interesting and have nanoparticles on them, so it is important to determine early on the nanoparticles of greatest interest.
In the case of nanoparticles, it is important for the microscopist to recognize that they are viewing an airy disc or &#8220;point spread function&#8221; of the object2. In general, the airy disc is useful in determining whether a plasmonic nanoparticle has a shape that is isotropic or anisotropic, but nanoparticle imaging is in fact much more complex than what is discussed here. Complex nanoparticle aggregates can sometimes resemble isotropic particles, and as a result, electron microscopy methods are then necessary to characterize the nanoparticle patterns2,16,18,19. To image plasmonic nanoparticles with a DIC microscope, it is crucial to use filtered imaging and to image the particles at one of their highly-absorbing plasmonic wavelengths6. Imaging at an improper wavelength or without filters can result in the capture of shadow-cast patterns that are difficult to decipher.
When imaging nanoparticles alongside objects that are larger than the diffraction limit of light, it is important to remember that the microscope&#8217;s objective &#8220;sees&#8221; a relatively flat focal plane. A common misconception of DIC is that it enables viewing of an object in actual 3D relief. This is caused by the shadow-cast patterning, which indeed makes many objects appear to be three-dimensional. However, to collect vertical information on multiple focal planes, it would be necessary to raise or lower the stage and collect a sequence of images. This can be very difficult to perform and to interpret, especially for thicker samples, such as cells. Thus, the microscopist needs a deep understanding of all materials involved when performing such experiments and must record the positions of the individual focal planes that were utilized.
Finally, the data analysis step is as critical as data collection. When measuring the contrast or intensity values of the sample, several factors should be kept in mind. Typically, the analyst is primarily interested in the minimum and maximum values for the particle of interest. When the contrast to noise ratio for the sample is sufficiently high, and if the background area is clean and evenly illuminated, then a simple geometric shape can be drawn around the sample region without concern of signal being introduced by contaminants. Furthermore, if the background is clean and evenly illuminated, a background measurement can be made in any area immediately adjacent to the sample. However, if there are contaminants or if the background is uneven, then the analyst must make a critical review of the sample&#8217;s environs, and the analyst needs to assess whether it is even possible to make a reasonable background measurement. It is also critical to measure the sample and background areas with the same-sized and shaped tool, in order to avoid the introduction of bias into the calculation. In general, smaller-sized measurement areas have a lower likelihood of detecting outliers (e.g., contaminants, bad pixels, etc.) but larger sampling areas often provide a more reliable measurement of the background&#8217;s mean value.

Since the 1980s, differential interference contrast (DIC) microscopy has largely been viewed as an important imaging method reserved for microscale objects within the biological sciences. However, during its development in the 1950s and 1960s, it was intended as a technique for materials science1. With the recent advancements in the material sciences related to plasmonic nanoparticles, an increased interest in the characterization of materials with optical microscopy has taken place.
Many optical techniques are certainly available for nanomaterial characterization (e.g., dark field, brightfield, polarized light, fluorescence, etc.). Dark field is widely popular in nanoparticle research, but it relies solely on the collection of scatter and provides limited information about complex samples2. Fluorescence can be useful, but only with samples that luminesce or that can be properly stained. DIC microscopy has several traits that make it a valuable tool for the analysis of nanoparticles. The most frequently stated advantages of DIC in comparison to other methods and in regards to plasmonic nanoparticles are: no sample staining required, no halo effects, shallow depth of field, and high lateral resolution3. DIC has additional strengths that are valuable to plasmonic nanoparticle research. First of all, two inherent and orthogonal polarization fields are present, and they can be measured simultaneously for spectroscopy purposes2. Secondly, the depolarized signal of nanoparticles is not captured in the final image2, which can be a cause for serious concern in dark field spectroscopy measurements.
The purpose of this article is to provide a clear methodology for utilizing transmitted-light Nomarski DIC microscopy to perform spectroscopy on plasmonic nanoparticles. Although DIC is a powerful technique that can be applied to highly diverse materials, it is also a technique that requires great skill and understanding to operate it properly when imaging nanoparticles. Transmission-based Nomarski DIC microscopy has a complex light path1 that will only be briefly reviewed here. The optical train of DIC is displayed in Figure 1. Light is transmitted through the microscope by first being passed through a polarizer and a beam-splitting Nomarski prism before being focused by the condenser onto the sample plane. After passing through the objective, the light encounters a beam-combining Nomarski prism and an analyzer before exiting to the detector. The two polarizers and Nomarski prisms are critical to formation of the DIC image and are responsible for producing DIC&#8217;s two orthogonal polarization fields1. For the reader interested in knowing more about the working principles and optical path of Nomarski DIC microscopes, or the differences between Nomarski DIC and other styles of DIC, please refer to other well-written accounts on these topics1,4,5,6,7.
It is equally important to understand the basic nature of plasmonic nanoparticles before attempting to perform spectroscopy on them, whether it be with Nomarski DIC, dark field, or any other microscopy technique. In the field of plasmonics, nanoparticles are defined as particulates with dimensions on the scale of 10-100 nm8,9. Nanoparticles can take on many shapes (e.g., spheres, rods, stars, dumbbells, etc.), and most of their important properties arise from interactions with light in the ultraviolet-visible-near infrared range of the electromagnetic spectrum. The term &#8220;plasmonic&#8221; is not restricted to nanoparticles10; however, when discussing nanoparticles, it is used in reference to localized surface plasmon resonance (LSPR). LSPR is a phenomenon in which the conduction electrons in a nanoparticle oscillate due to a Coulombic interaction with electromagnetic radiation of a highly specific and relatively narrow frequency band8. At these same frequencies, plasmonic nanoparticles exhibit increased absorption and scattering of light, making them observable with optical microscopy. In many cases, it is preferred to observe the nanoparticles while placing bandpass filters before the condenser2, to improve imaging contrast and to eliminate light that fails to induce the LSPR effect. Using filters also makes it possible to perform single particle spectroscopy experiments.
LSPR-related optical behavior is highly dependent on the size and shape of the nanoparticles, and it can be investigated with many optical microscopy techniques. However, in order to decipher orientation information of plasmonic nanoparticles with an anisotropic (i.e., non-spherical) shape, it is necessary to utilize polarization of the light field. By carefully rotating the polarization field or the sample substrate at small increments, it is possible to monitor the orientation-dependent spectroscopic properties of individual nanoparticles. Rotation and polarization can also aid in determining whether a spectral feature is due to a dipolar or higher order oscillation of the nanoparticle&#8217;s surface electrons. However, in the case of isotropic (i.e., spherical) nanoparticles, the spectral profile remains essentially unchanged upon rotating the sample under polarized light.
When viewed through a DIC microscope (Figure 2), nanoparticles have an airy disk with a shadow-cast white-and-black appearance against a gray background. Spherical nanoparticles will retain this appearance under rotation and with the changing of bandpass filters; however, the particles will gradually fade from view as the filter&#8217;s central wavelength becomes further separated from the sphere&#8217;s only dipolar LSPR wavelength11. The appearance of nanorods can change quite dramatically as they are rotated2. Nanorods have two LSPR bands with dipolar behavior, the location of which are based on the physical dimensions of the nanorods. When the longitudinal axis of a nanorod is oriented parallel to one of the DIC polarization fields, the airy disc will appear all white or all black if viewed with a bandpass filter associated with that LSPR wavelength. After rotating the sample 90&#176;, it will take on the opposite color. Alternatively, since the transverse axis of a nanorod is perpendicular to the longitudinal axis, the rod will take on the opposite color when switching between filters that match the LSPR wavelengths for the two axes. At other orientations and filter settings, nanorods will appear more like spheres, presenting a variety of shadow-cast airy disc patterns. For nanorods with a transverse axis &#60; 25 nm, it can be difficult to detect signal at that LSPR&#8217;s wavelength using DIC microscopy.
To perform single particle spectroscopy, it is important to use the correct optical components and to align them properly. An objective capable of DIC microscopy must be used. For single particle experiments, 80x or 100x oil objectives are ideal. Nomarski DIC prisms ordinarily come in three varieties: standard, high contrast, and high resolution. The ideal type highly depends on the purpose of the experiment and the size of the nanoparticles. Standard prisms are fine for many experiments; but when working with smaller nanoparticles (&#60; 50 nm), high contrast prisms can be beneficial, since particle contrast decreases as the particles decrease in size11. Adjusting the DIC contrast is achieved either by rotating a polarizer or by translating one of the DIC prisms, depending on the microscope brand or model6.
After setting Kohler illumination and the polarizer settings, it is critical to not readjust these settings while collecting spectroscopy data. Furthermore, a constant average background signal must be maintained at all times during data collection, even when switching between filters and angle settings. The actual ideal background value depends on the dynamic range of the scientific camera, but in general, the background should be in the range of 15%&#8211;40% of the maximum detection level of the camera. This reduces the likelihood of saturating the camera sensor while enabling optimal particle contrast. For collecting spectroscopy data, it is necessary to work with a scientific camera that captures images in black and white, as opposed to a color camera.
Sample preparation is another critical aspect of imaging plasmonic nanoparticles. It is imperative that operators of DIC microscopy have an understanding of the sample&#8217;s optical properties and of the sample&#8217;s substrate. &#8220;Pre-cleaned&#8221; microscope glass is not sufficiently prepared for imaging nanoparticles, and it must be properly re-cleaned before sample deposition to ensure unobstructed observation of the sample. Many cleaning protocols for microscope slides have been previously documented12, but it is not a step that is normally reported in experimental studies.
Finally, data analysis methods are the final component to single particle spectroscopy. The maximum and minimum intensities for each nanoparticle must be measured, as well as the local background average. Particles of interest should be located in areas with no background debris, substrate defects, or uneven illumination. One method for determining the spectral profile of a nanoparticle is by calculating particle contrast at each wavelength, using the equation below11,13,14,15:
<img alt="Equation" src="/files/ftp_upload/59411/59411eq1.jpg" />
Alternatively, a single particle&#8217;s spectrum can be split into its individual maximum and minimum signal components, which represent DIC&#8217;s two polarization fields, thereby displaying the two simultaneously-collected directionally-dependent spectra, through the two equations:
<img alt="Equation" src="/files/ftp_upload/59411/59411eq2.jpg" />
<img alt="Equation" src="/files/ftp_upload/59411/59411eq3.jpg" />

<table><tbody><tr><td>Contrad 70</td><td>Decon Labs, Inc.</td><td>1002</td><td>For cleaning microscope glass, Available through many chemical suppliers</td></tr><tr><td>Ethanol</td><td>Fisher Scientific</td><td>A962-4</td><td>For cleaning and storing microscope glass</td></tr><tr><td>Glass microscope cover slips</td><td>Ted Pella</td><td>260148</td><td></td></tr><tr><td>Glass microscope slides</td><td>Ted Pella</td><td>26007</td><td></td></tr><tr><td>Gold nanorods</td><td>Nanopartz</td><td>DIAM-SPR-25-650</td><td></td></tr><tr><td>Gold nanospheres (80 nm)</td><td>Sigma Aldrich</td><td>742023-25ML</td><td></td></tr><tr><td>ImageJ</td><td>NIH</td><td>N/A</td><td>Free Software availabe for data analysis from NIJ</td></tr><tr><td>Nail polish</td><td>Electron Microscopy Sciences</td><td>72180</td><td></td></tr><tr><td>Nikon Ti-E microscope</td><td>Nikon</td><td>N/A</td><td></td></tr><tr><td>Nitrogen gas</td><td>Airgas</td><td>N/A</td><td></td></tr><tr><td>ORCA Flash 4.0 V2+ digital sCMOS camera</td><td>Hamamatsu</td><td>77054098</td><td></td></tr><tr><td>Scribing pen</td><td>Amazon</td><td>N/A</td><td>Many options available online for under $10. Not necessary to buy an expensive version.</td></tr><tr><td>Ultrapure water</td><td></td><td></td><td>18 megaohm</td></tr></tbody></table>

performing spectroscopy on plasmonic nanoparticles with transmission-based nomarski-type differential interference contrast microscopy

Differential interference contrast (DIC) microscopy is a powerful imaging tool that is most commonly employed for imaging microscale objects using visible-range light. The purpose of this protocol is to detail a proven method for preparing plasmonic nanoparticle samples and performing single particle spectroscopy on them with DIC microscopy. Several important steps must be followed carefully in order to perform repeatable spectroscopy experiments. First, landmarks can be etched into the sample substrate, which aids in locating the sample surface and in tracking the region of interest during experiments. Next, the substrate must be properly cleaned of debris and contaminants that can otherwise hinder or obscure examination of the sample. Once a sample is properly prepared, the optical path of the microscope must be aligned, using Kohler Illumination. With a standard Nomarski style DIC microscope, rotation of the sample may be necessary, particularly when the plasmonic nanoparticles exhibit orientation-dependent optical properties. Because DIC microscopy has two inherent orthogonal polarization fields, the wavelength-dependent DIC contrast pattern reveals the orientation of rod-shaped plasmonic nanoparticles. Finally, data acquisition and data analyses must be carefully performed. It is common to represent DIC-based spectroscopy data as a contrast value, but it is also possible to present it as intensity data. In this demonstration of DIC for single particle spectroscopy, the focus is on spherical and rod-shaped gold nanoparticles.

When working with samples that are large enough to be seen with the naked eye, placing landmarks on the glass substrate is not normally required. However, when working with nanomaterials or when rotation of the sample is required, landmarks can provide an easy method for locating, distinguishing, and tracking the orientation of the sample. Although more sophisticated techniques can be utilized for leaving landmarks on glass substrates17, scratching the glass with a scribing pen is an economical and simple method that works in many situations. It is important to avoid examining sample regions that are immediately adjacent to these landmarks, since scratch marks create a complex background with the potential of impacting data (Figure 3). However, at the tips of the scratch marks, &#8220;spider webs&#8221; often extend outward from the scratch. These lines are quite valuable as landmarks, but again, nanoparticles should&#160;be avoided if&#160;they overlap with these defects.
In order to achieve optimal imaging with DIC microscopy, it is of vital importance to determine the proper focal plane. Objects that are slightly out of focus will appear fuzzy, have blurred edges, and have decreased contrast. Figure 4 displays gold nanoparticles that are out of focus to varying degrees. Nanoparticles in the bottom right corner are in focus, while nanoparticles become farther out of focus as they approach the upper left corner of this image. Because DIC has a shallow depth of field, it is not uncommon for some nanoparticles to be in focus while others are out of focus when imaging them on a glass substrate. As a result, it is critical to consistently focus on the same exact particles when making adjustments to the microscope during an experiment.
Figure 5 provides an example of the effect of adjusting the polarizer settings while imaging gold nanospheres. Five nanoparticles are in focus, while one is slightly out of focus. A 540 nm bandpass filter with 10 nm FWHM was also in the optical path. In this series of images, the background brightness was adjusted with ImageJ after image acquisition in order to make the five particles more apparent against the background. When the polarizer is set at 0&#176; in a de Senarmont designed Nomarski DIC microscope, it is orthogonal to the analyzer (Figure 5A). At 0&#176;, the particles appear mostly white, with a dark stripe running across their mid-section. This is indicative of cross-polarization for nanosphere samples. When the polarizer is rotated to different angles (Figure 5B-E), the particles appear to be casting dark shadows towards the southwest. The black and white components to the signal arise as a result of DIC&#8217;s two polarization fields and provide information about the orientation of plasmonic nanoparticles when working with bandpass filters. As the polarizer is rotated towards higher angles, the shadow pattern remains similar. However, the particle contrast values change dramatically. This is best demonstrated by measuring the contrast values for the individual particles, using the equation provided above. The particle highlighted with the black box has contrast values of 0.65 (crossed polarizers), 0.84 (polarizer shift of 5&#176;), 1.10 (10&#176;), 0.44 (20&#176;), and 0.23 (45&#176;). Therefore, for this sample, the optimal imaging setting is with a polarizer shift of 10&#176;. Plasmonic nanoparticles often require a polarizer setting in the range of 5&#176;&#8211;15&#176;, and smaller increments than these should normally be used to identify the ideal setting. For further information on the imaging and analysis of spherical gold nanoparticles, readers are referred to the prior work by Sun et al.11.
Anisotropic-shaped nanoparticles produce patterns of higher complexity than nanospheres. Gold nanorods were imaged (Figure 6) at their longitudinal LSPR wavelength, 650 nm. In the initial image (Figure 6A), five bright nanorods and several dimmer particles are apparent. Instead of having a shadow-cast appearance, three of the rods have predominately black airy discs while two are mostly white. The polarizer was set at 10&#176; to the left of the crossed polarization setting. In Figure 6B, crossed polarization was used; only three of the particles appear, as fully white airy discs. The others have disappeared or appear to be slightly out of focus. With the polarizer set at 10&#176; to the right of crossed polarization (Figure 6C), the patterns are now reversed of what was observed in Figure 6A. The polarizer was next turned to 45&#176; right of crossed polarization (Figure 6D), the maximum setting, to demonstrate that particles retain their colors at this setting, but contrast has declined significantly. In the remaining figure panels, the collection of nanorods was incrementally rotated a full 90&#176; clockwise while the polarizer was set at 10&#176; to the right of crossed polarization. The pattern gradually changes for each nanorod, and after a full 90&#176; rotation, the particles have reversed their colors from the initial setting. In brief, if one of the axes of a plasmonic nanorod is lined up with one of the two polarization fields, and if the nanorod is imaged at that axis&#8217; LSPR wavelength, the nanorod will appear to be mostly white or mostly black, depending on which polarization field it is aligned with (Figure 6A,C)2. If the nanoparticle is rotated a full 90&#176; (Figure 6H), it will now be lined up with the opposite polarization field and take on the opposite color. If instead the nanoparticle was rotated only 45&#176; (Figure 6F), then it will be in a position where the particle will exhibit its greatest shadow-cast appearance, showing striking similarity to what is observed with the plasmonic nanospheres. As a result of this optical behavior, plasmonic nanoparticles with an anisotropic shape often look flat instead of having the three-dimensional shadow-cast appearance of nanospheres. The result of this difference in optical behavior is that it can be exploited in order to distinguish between plasmonic nanoparticles that are spherical and anisotropic in shape, as has been previously discussed in multiple research studies2,3,6,7,11,13,16.
Finally, Figure 7 displays representative single particle spectroscopy data, as contrast of a gold nanosphere (Figure 7A)11, intensity of a single gold nanorod with its longitudinal axis oriented parallel to one of the polarization fields (Figure 7B)6, and intensity profile of a single gold nanorod at its LSPR wavelength and during rotation of the stage (Figure 7C)6. Either method of presentation reveals the width and location of the LSPR effect. For plasmonic nanoparticles with an anisotropic shape, the intensity and rotation data reveal the directionality of the effect, and hence, the orientation of the particle on the sample substrate, which has been previously proven through correlative studies on such particles using DIC and transmission electron microscopy2,16,18.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59411/59411fig1.jpg" /> 
Figure 1: The light path in transmitted-light Nomarski-based DIC microscopy. After leaving the light source (S), light passes through a polarizer (P), a beam-shearing Nomarski prism (NP), the condenser (C), the focal plane (FP), the objective (O), a beam-combining Nomarski prism (NP), the analyzer (A), and finally the detector (D). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59411/59411fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59411/59411fig2.jpg" /> 
Figure 2: Examples of plasmonic nanoparticles imaged at their LSPR wavelengths with 10 nm bandpass filters, using a DIC microscope. Both images are collected at 100x. (A) Silver nanospheres with 40 nm diameter imaged at 480 nm with a bandpass filter having 10 nm FWHM. (B) Rod-like gold nanoparticles imaged at 700 nm using a bandpass filter with 10 nm FWHM. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59411/59411fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59411/59411fig3.jpg" /> 
Figure 3: Image of a scratch made into a glass cover slip with a scribing pen. Near the end of the actual indentation, a series of narrow and shallow &#8220;spider web&#8221; lines branch out from the scratch itself, resulting in a pattern that can be utilized as an imaging landmark. This image was collected using 100x magnification and broadband white light. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59411/59411fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59411/59411fig4.jpg" /> 
Figure 4: Gold nanospheres imaged with broadband white light at 100x. Particles in the lower right are in focus but particles drift farther from the focal plane towards the upper left corner. Object in middle of image is debris. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59411/59411fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59411/59411fig5.jpg" /> 
Figure 5: Gold nanospheres imaged under a series of different polarizer settings at a wavelength of 540 nm and a magnification of 100x. Background brightness was adjusted post-imaging with ImageJ to make particles more apparent. Polarizer setting of (A) 0&#176; (polarizer orthogonal to analyzer), (B) 5&#176;, (C) 10&#176; (the best contrast of this series of images), (D) 20&#176;, and (E) 45&#176;. Measured contrast of particle in black box is (A) 0.65, (B) 0.84, (C) 1.10, (D) 0.44, and (E) 0.23. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59411/59411fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59411/59411fig6.jpg" /> 
Figure 6: Example of imaging anisotropic plasmonic nanoparticles: gold nanorods at their longitudinal LSPR wavelength of 650 nm and a magnification of 100x. Particles of main interest are enclosed in yellow box. Polarizer settings are: (A) left 10&#176;, (B) 0&#176;, (C) right 10&#176;, (D) right 45&#176;. With the polarizer set to right 10&#176;, the stage was rotated clockwise by (E) 20&#176;, (F) 45&#176;, (G) 70&#176;, and (H) 90&#176;. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59411/59411fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/59411/59411fig7.jpg" /> 
Figure 7: Representative results of single particle spectroscopy data. (A) Gold nanosphere spectroscopy displayed in terms of DIC Contrast. Each data point represents an average of 20 nanospheres for each particle diameter, and data capture relied on 10nm FWHM bandpass filters. (B) A single gold nanorod displayed as DIC Intensity data, using two different polarizer settings (2&#176; on either side of crossed polarization). (C) DIC Intensity data for a single gold nanorod at the LSPR wavelength of 680 nm, while it was rotated 180&#176; and the polarizer was held at 2&#176; off the crossed polarization position. Figure 7A is adapted with permission from Sun et al., Analytical Chemistry. 81 (22), 9203-9208 (2009), and Figure 7B,C from Stender et al., Analytical Chemistry. 84 (12), 5210-5215 (2012). Copyright American Chemical Society. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/59411/59411fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Performing Spectroscopy on Plasmonic Nanoparticles with Transmission-Based Nomarski-Type Differential Interference Contrast Microscopy at JoVE.com

Performing Spectroscopy on Plasmonic Nanoparticles with Transmission-Based Nomarski-Type Differential Interference Contrast Microscopy

The goal of this protocol is to detail a proven approach for the preparation of plasmonic nanoparticle samples and for performing single particle spectroscopy on them with differential interference contrast (DIC) microscopy.

Differential interference contrast (DIC) microscopy is a powerful imaging tool that is most commonly employed for imaging microscale objects using ...

performing-spectroscopy-on-plasmonic-nanoparticles-with-transmission

Nanyang Technological University

Research

JoVE Journal

Chemistry

7.5K Views.  Ohio University. The goal of this protocol is to detail a proven approach for the preparation of plasmonic nanoparticle samples and for performing single particle spectroscopy on them with differential interference contrast (DIC) microscopy.

Video: Performing Spectroscopy on Plasmonic Nanoparticles with Transmission-Based Nomarski-Type Differential Interference Contrast Microscopy

Evaluating Plasmonic Transport in Current-carrying Silver Nanowires

Plasmonics is an emerging technology capable of simultaneously transporting a plasmonic signal and an electronic signal on the same information support1,2,3. In this context, metal nanowires are especially desirable for realizing dense routing networks4. A prerequisite to operate such shared nanowire-based platform relies on our ability to electrically contact individual metal nanowires and efficiently excite surface plasmon polaritons5 in this information support. In this article, we describe a protocol to bring electrical terminals to chemically-synthesized silver nanowires6 randomly distributed on a glass substrate7. The positions of the nanowire ends with respect to predefined landmarks are precisely located using standard optical transmission microscopy before encapsulation in an electron-sensitive resist. Trenches representing the electrode layout are subsequently designed by electron-beam lithography. Metal electrodes are then fabricated by thermally evaporating a Cr/Au layer followed by a chemical lift-off. The contacted silver nanowires are finally transferred to a leakage radiation microscope for surface plasmon excitation and characterization8,9. Surface plasmons are launched in the nanowires by focusing a near infrared laser beam on a diffraction-limited spot overlapping one nanowire extremity5,9. For sufficiently large nanowires, the surface plasmon mode leaks into the glass substrate9,10. This leakage radiation is readily detected, imaged, and analyzed in the different conjugate planes in leakage radiation microscopy9,11. The electrical terminals do not affect the plasmon propagation. However, a current-induced morphological deterioration of the nanowire drastically degrades the flow of surface plasmons. The combination of surface plasmon leakage radiation microscopy with a simultaneous analysis of the nanowire electrical transport characteristics reveals the intrinsic limitations of such plasmonic circuitry.

Silver nanowires can simultaneously transport electrons and optical information in the form of surface plasmons. A procedure is described here to realize such a shared circuitry and the limitations at propagating both information carriers are evaluated.

Plasmonics is an emerging technology capable of simultaneously transporting a plasmonic signal and an electronic signal on the same information ...

Engineering

Utilization of Plasmonic and Photonic Crystal Nanostructures for Enhanced Micro- and Nanoparticle Manipulation

A method to manipulate the position and orientation of submicron particles nondestructively would be an incredibly useful tool for basic biological research. Perhaps the most widely used physical force to achieve noninvasive manipulation of small particles has been dielectrophoresis(DEP).1 However, DEP on its own lacks the versatility and precision that are desired when manipulating cells since it is traditionally done with stationary electrodes. Optical tweezers, which utilize a three dimensional electromagnetic field gradient to exert forces on small particles, achieve this desired versatility and precision.2 However, a major drawback of this approach is the high radiation intensity required to achieve the necessary force to trap a particle which can damage biological samples.3 A solution that allows trapping and sorting with lower optical intensities are optoelectronic tweezers (OET) but OET's have limitations with fine manipulation of small particles; being DEP-based technology also puts constraint on the property of the solution.4,5
This video article will describe two methods that decrease the intensity of the radiation needed for optical manipulation of living cells and also describe a method for orientation control. The first method is plasmonic tweezers which use a random gold nanoparticle (AuNP) array as a substrate for the sample as shown in Figure 1. The AuNP array converts the incident photons into localized surface plasmons (LSP) which consist of resonant dipole moments that radiate and generate a patterned radiation field with a large gradient in the cell solution. Initial work on surface plasmon enhanced trapping by Righini et al and our own modeling have shown the fields generated by the plasmonic substrate reduce the initial intensity required by enhancing the gradient field that traps the particle.6,7,8 The plasmonic approach allows for fine orientation control of ellipsoidal particles and cells with low optical intensities because of more efficient optical energy conversion into mechanical energy and a dipole-dependent radiation field. These fields are shown in figure 2 and the low trapping intensities are detailed in figures 4 and 5. The main problems with plasmonic tweezers are that the LSP's generate a considerable amount of heat and the trapping is only two dimensional. This heat generates convective flows and thermophoresis which can be powerful enough to expel submicron particles from the trap.9,10 The second approach that we will describe is utilizing periodic dielectric nanostructures to scatter incident light very efficiently into diffraction modes, as shown in figure 6.11 Ideally, one would make this structure out of a dielectric material to avoid the same heating problems experienced with the plasmonic tweezers but in our approach an aluminum-coated diffraction grating is used as a one-dimensional periodic dielectric nanostructure. Although it is not a semiconductor, it did not experience significant heating and effectively trapped small particles with low trapping intensities, as shown in figure 7. Alignment of particles with the grating substrate conceptually validates the proposition that a 2-D photonic crystal could allow precise rotation of non-spherical micron sized particles.10 The efficiencies of these optical traps are increased due to the enhanced fields produced by the nanostructures described in this paper.

Plasmonic tweezers and photonic crystal nanostructures are shown to produce useful enhancements in the efficiency and orientation control of optically trapping micro- and nano-particles.

A method to manipulate the position and orientation of submicron particles nondestructively would be an incredibly useful tool for basic biological ...

Bioengineering

Analyzing the Movement of the Nauplius 'Artemia salina' by Optical Tracking of Plasmonic Nanoparticles

We demonstrate how optical tweezers may provide a sensitive tool to analyze the fluidic vibrations generated by the movement of small aquatic organisms. A single gold nanoparticle held by an optical tweezer is used as a sensor to quantify the rhythmic motion of a Nauplius larva (Artemia salina) in a water sample. This is achieved by monitoring the time dependent displacement of the trapped nanoparticle as a consequence of the Nauplius activity. A Fourier analysis of the nanoparticle&#39;s position then yields a frequency spectrum that is characteristic to the motion of the observed species. This experiment demonstrates the capability of this method to measure and characterize the activity of small aquatic larvae without the requirement to observe them directly and to gain information about the position of the larvae with respect to the trapped particle. Overall, this approach could give an insight on the vitality of certain species found in an aquatic ecosystem and could expand the range of conventional methods for analyzing water samples.

Analyzing the Movement of the Nauplius 'Artemia salina' by Optical Tracking of Plasmonic Nanoparticles

We use optical tracking of plasmonic nanoparticles to probe and characterize the frequency movements of aquatic organisms.

We demonstrate how optical tweezers may provide a sensitive tool to analyze the fluidic vibrations generated by the movement of small aquatic organisms. A ...

Measurement of Scattering Nonlinearities from a Single Plasmonic Nanoparticle

Plasmonics, which are based on the collective oscillation of electrons due to light excitation, involve strongly enhanced local electric fields and thus have potential applications in nonlinear optics, which requires extraordinary optical intensity. One of the most studied nonlinearities in plasmonics is nonlinear absorption, including saturation and reverse saturation behaviors. Although scattering and absorption in nanoparticles are closely correlated by the Mie theory, there has been no report of nonlinearities in plasmonic scattering until very recently. 
Last year, not only saturation, but also reverse saturation of scattering in an isolated plasmonic particle was demonstrated for the first time. The results showed that saturable scattering exhibits clear wavelength dependence, which seems to be directly linked to the localized surface plasmon resonance (LSPR). Combined with the intensity-dependent measurements, the results suggest the possibility of a common mechanism underlying the nonlinear behaviors of scattering and absorption. These nonlinearities of scattering from a single gold nanosphere (GNS) are widely applicable, including in super-resolution microscopy and optical switches. 
In this paper, it is described in detail how to measure nonlinearity of scattering in a single GNP and how to employ the super-resolution technique to enhance the optical imaging resolution based on saturable scattering. This discovery features the first super-resolution microscopy based on nonlinear scattering, which is a novel non-bleaching contrast method that can achieve a resolution as low as l/8 and will potentially be useful in biomedicine and material studies.

Saturable and reverse saturable scattering were discovered in isolated plasmonic particles and adopted as a novel non-bleaching contrast method in super-resolution microscopy. Here the experimental procedures of detecting and extracting nonlinear scattering are explained in detail, as well as how to enhance resolution with the aid of saturated excitation microscopy.

Plasmonics, which are based on the collective oscillation of electrons due to light excitation, involve strongly enhanced local electric fields and thus ...

Colloidal Synthesis of Nanopatch Antennas for Applications in Plasmonics and Nanophotonics

We present a method for colloidal synthesis of silver nanocubes and the use of these in combination with a smooth gold film, to fabricate plasmonic nanoscale patch antennas. This includes a detailed procedure for the fabrication of thin films with a well-controlled thickness over macroscopic areas using layer-by-layer deposition of polyelectrolyte polymers, namely poly(allylamine) hydrochloride (PAH) and polystyrene sulfonate (PSS). These polyelectrolyte spacer layers serve as a dielectric gap in between silver nanocubes and a gold film. By controlling the size of the nanocubes or the gap thickness, the plasmon resonance can be tuned from about 500 nm to 700 nm. Next, we demonstrate how to incorporate organic sulfo-cyanine5 carboxylic acid (Cy5) dye molecules into the dielectric polymer gap region of the nanopatch antennas. Finally, we show greatly enhanced fluorescence of the Cy5 dyes by spectrally matching the plasmon resonance with the excitation energy and the Cy5 absorption peak. The method presented here enables the fabrication of plasmonic nanopatch antennas with well-controlled dimensions utilizing colloidal synthesis and a layer-by-layer dip-coating process with the potential for low cost and large-scale production. These nanopatch antennas hold great promise for practical applications, for example in sensing, ultrafast optoelectronic devices and for high-efficiency photodetectors.

A protocol for the colloidal synthesis of silver nanocubes and fabrication of plasmonic nanoscale patch antennas with sub-10 nm gaps is presented.

We present a method for colloidal synthesis of silver nanocubes and the use of these in combination with a smooth gold film, to fabricate plasmonic ...

Plasmonic Trapping and Release of Nanoparticles in a Monitoring Environment

Plasmonic tweezers use surface plasmon polaritons to confine polarizable nanoscale objects. Among the various designs of plasmonic tweezers, only a few can observe immobilized particles. Moreover, a limited number of studies have experimentally measured the exertable forces on the particles. The designs can be classified as the protruding nanodisk type or the suppressed nanohole type. For the latter, microscopic observation is extremely challenging. In this paper, a new plasmonic tweezer system is introduced to monitor particles, both in directions parallel and orthogonal to the symmetric axis of a plasmonic nanohole structure. This feature enables us to observe the movement of each particle near the rim of the nanohole. Furthermore, we can quantitatively estimate the maximal trapping forces using a new fluidic channel.

A microchip fabrication process that incorporates plasmonic tweezers is presented here. The microchip enables the imaging of a trapped particle to measure maximal trapping forces.

Plasmonic tweezers use surface plasmon polaritons to confine polarizable nanoscale objects. Among the various designs of plasmonic tweezers, only a few ...

Visualizing Diffusional Dynamics of Gold Nanorods on Cell Membrane using Single Nanoparticle Darkfield Microscopy

Analyzing the diffusional dynamics of nanoparticles on cell membrane plays a significant role in better understanding the cellular uptake process and provides a theoretical basis for the rational design of nano-medicine delivery. Single particle tracking (SPT) analysis could probe the position and orientation of individual nanoparticles on cell membrane, and reveal their translational and rotational states. Here, we show how to use traditional dark-field microscopy to monitor the dynamics of gold nanorods (AuNRs) on live cell membrane. We also show how to extract the location and orientation of AuNRs using ImageJ and MATLAB, and how to characterize the diffusive states of AuNRs. Statistical analysis of hundreds of particles show that single AuNRs perform Brownian motion on the surface of U87 MG cell membrane. However, individual long trajectory analysis shows that AuNRs have two distinctly different types of motion states on the membrane, namely long-range transport and limited-area confinement. Our SPT methods can be potentially used to study the surface or intracellular particle diffusion in different biological cells and can become a powerful tool for investigations of complex cellular mechanisms.

Here, we show the use of traditional dark-field microscopy to monitor the dynamics of gold nanorods (AuNRs) on cell membrane. The location and orientation of single AuNRs are detected using ImageJ and MATLAB, and the diffusive states of AuNRs are characterized by single particle tracking analysis.

Analyzing the diffusional dynamics of nanoparticles on cell membrane plays a significant role in better understanding the cellular uptake process and ...

Biochemistry

Spectral and Angle-Resolved Magneto-Optical Characterization of Photonic Nanostructures

Photonic crystals are periodic nanostructures that can support a variety of confined electromagnetic modes. Such confined modes are usually accompanied by local enhancement of electric field intensity that strengthens light-matter interactions, enabling applications such as surface-enhanced Raman scattering (SERS) and surface plasmon enhanced sensing. In the presence of magneto-optically active materials, the local field enhancement gives rise to anomalous magneto-optical activity. Typically, the confined modes of a given photonic crystal depend strongly on the wavelength and incidence angle of the incident electromagnetic radiation. Thus, spectral and angular-resolved measurements are needed to fully identify them as well as to establish their relationship with the magneto-optical activity of the crystal. In this article, we describe how to use a Fourier-plane (back focal plane) microscope to characterize magneto-optically active samples. As a model system, here we use a plasmonic grating built out of magneto-optically active Au/Co/Au multilayer. In the experiments, we apply a magnetic field on the grating in situ and measure its reciprocal space response, obtaining the magneto-optical response of the grating over a range of wavelengths and incident angles. This information enables us to build a complete map of the plasmonic band structure of the grating and the angle and wavelength dependent magneto-optical activity. These two images allow us to pinpoint the effect that the plasmon resonances have on the magneto-optical response of the grating. The relatively small magnitude of magneto-optical effects requires a careful treatment of the acquired optical signals. To this end, an image processing protocol for obtaining magneto-optical response from the acquired raw data is laid out.

Photonic band structure enables understanding how confined electromagnetic modes propagate within a photonic crystal. In photonic crystals that incorporate magnetic elements, such confined and resonant optical modes are accompanied by enhanced and modified magneto-optical activity. We describe a measurement procedure to extract the magneto-optical band structure by Fourier space microscopy.

Photonic crystals are periodic nanostructures that can support a variety of confined electromagnetic modes. Such confined modes are usually accompanied by ...

Monitoring Conformational Dynamics of Single Unmodified Proteins using Plasmonic Nanotweezers

Current single-molecule techniques to characterize proteins typically require labels, tethers, or the use of non-native solution conditions. Such changes can alter protein biophysics and reduce the usefulness of the data acquired. Plasmonic nanotweezers is a technique that uses localized surface plasmon resonance (LSPR) on gold nanostructures to enhance the electric field within a confined hotspot region. This field enhancement allows for the use of low laser powers to trap single nanoparticles far smaller than conventional optical tweezers, down to only a few nanometers in diameter, such as single proteins. Trapping of single protein molecules within the hotspot region induces a shift in the local refractive index (nprotein &#62; nwater), altering light scattering as a product of the molecule&#39;s polarisability, which is affected by its volume, shape anisotropy, and refractive index. An avalanche photodiode (APD) collects the subsequent changes in light scattering. These alterations can then be analyzed to determine changes in the trapped molecule, including its size, global conformation, and dynamics of conformational change over time. The incorporation of microfluidics within the system allows for controlled environmental changes and real-time monitoring of their subsequent effects on the molecule. In this protocol, we demonstrate the steps to trap single protein molecules, alter their environmental solution conditions, and monitor their corresponding conformational changes using a plasmonic nanotweezers system.

Plasmonic nanotweezers use localized surface plasmon resonance in gold nanostructures to trap single nanoparticles, including proteins, within a nanometer-scale optical field. Changes in the scattered signal reveal protein presence and conformational dynamics, enabling monitoring without fluorophore modifications or surface tethering.

Current single-molecule techniques to characterize proteins typically require labels, tethers, or the use of non-native solution conditions. Such changes ...

Optical Trapping of Plasmonic Nanoparticles for In Situ Surface-Enhanced Raman Spectroscopy Characterizations

Surface-enhanced Raman spectroscopy (SERS) enables the ultrasensitive detection of analyte molecules in various applications due to the enhanced electric field of metallic nanostructures. Salt-induced silver nanoparticle aggregation is the most popular method for generating SERS-active substrates; however, it is limited by poor reproducibility, stability, and biocompatibility. The present protocol integrates optical manipulation and SERS detection to develop an efficient analytical platform to address this. A 1064 nm trapping laser and a 532 nm Raman probe laser are combined in a microscope to assemble silver nanoparticles, which generate plasmonic hotspots for in situ SERS measurements in aqueous environments. Without aggregating agents, this dynamic plasmonic silver nanoparticle assembly enables an approximately 50-fold enhancement of the analyte molecule signal. Moreover, it provides spatial and temporal control to form the SERS-active assembly in as low as 0.05 nM analyte-coated silver nanoparticle solution, which minimizes the potential perturbation for in vivo analysis. Hence, this optical trapping-integrated SERS platform holds great potential for efficient, reproducible, and stable molecular analyses in liquids, especially in aqueous physiological environments.

Optical Trapping of Plasmonic Nanoparticles for In Situ Surface-Enhanced Raman Spectroscopy Characterizations

The present protocol describes a convenient approach to integrating optical trapping and surface-enhanced Raman spectroscopy (SERS) to manipulate plasmonic nanoparticles for sensitive molecular detection. Without aggregating agents, the trapping laser assembles plasmonic nanoparticles to enhance the SERS signals of target analytes for in situ spectroscopic measurements.

Surface-enhanced Raman spectroscopy (SERS) enables the ultrasensitive detection of analyte molecules in various applications due to the enhanced electric ...

Performing Spectroscopy on Plasmonic Nanoparticles with Transmission-Based Nomarski-Type Differential Interference Contrast Microscopy

Summary

Explore More Videos

Chapters in this video

Evaluating Plasmonic Transport in Current-carrying Silver Nanowires

Utilization of Plasmonic and Photonic Crystal Nanostructures for Enhanced Micro- and Nanoparticle Manipulation

Analyzing the Movement of the Nauplius 'Artemia salina' by Optical Tracking of Plasmonic Nanoparticles

Measurement of Scattering Nonlinearities from a Single Plasmonic Nanoparticle

Colloidal Synthesis of Nanopatch Antennas for Applications in Plasmonics and Nanophotonics

Plasmonic Trapping and Release of Nanoparticles in a Monitoring Environment

Visualizing Diffusional Dynamics of Gold Nanorods on Cell Membrane using Single Nanoparticle Darkfield Microscopy

Spectral and Angle-Resolved Magneto-Optical Characterization of Photonic Nanostructures

Monitoring Conformational Dynamics of Single Unmodified Proteins using Plasmonic Nanotweezers

Optical Trapping of Plasmonic Nanoparticles for In Situ Surface-Enhanced Raman Spectroscopy Characterizations