This research was supported by EPSRC Grants: Raman Nanotheranostics (EP/R020965/1) and CONTRAST facility (EP/S009957/1).

1. Switching on the laser system
<ol>
	<li>Switch on the interlock system and select arm laser before starting the system.</li>
	<li>Turn on the PC with the software to control the dual-output femtosecond laser.</li>
	<li>Load the software for the dual-output femtosecond laser; this software enables the laser to be powered on and off and directly controls the wavelength of the pump beam.</li>
	<li>Switch on the laser emission by holding down on the Power icon for a count of 3.</li>
	<li>Wait until the laser has warmed up and the laser-ready light is lit green in the software.</li>
	<li>The wavelength of the pump beam is directly controlled through the software; this can range from 680-1,300 nm. For cellular imaging in the C-H Raman vibrational region, select 802 nm.</li>
	<li>Open the shutters of the tunable pump beam and 1,045 nm Stokes beam by holding down on the aperture images for 3 s. 
	CAUTION: Laser safety guidance the dual-output femtosecond laser is a class IV infra-red pulsed laser and can cause permanent damage to eyesight. Therefore, all laser safety local rules must be followed while carrying out this procedure: (i) Only authorized users who have received laser safety training can carry out the procedure. (ii) Entry to the lab is&#160;via&#160;swipe card access with an interlock on the room door. (iii) For most of the operation, the laser beams are fully enclosed. (iv) When the laser beam is exposed, laser safety goggles must be worn (use the LG9 laser safety goggles, which give OD 7+ blocking of both the pump and Stokes beams).</li>
</ol>
2. Switching on the Spectral Focusing Timing and Recombination Unit (SF-TRU)
<ol>
	<li>Load the ATM software on the same PC with the laser software, which controls the delay stages and laser attenuators in the SF-TRU unit. 
	NOTE: This unit is responsible for making sure the pump and Stokes beams are spatially overlapped, controlling the dispersion in the pump and Stokes beams, and controlling the time delay between the pump and Stokes beams.The polarization of both pump and Stokes beams is vertical. Note that each beam is equipped with a half-wave plate to independently control polarizations before exiting the SF-TRU.</li>
	<li>Ensure that both the pump and Stokes lasers are attenuated to &#60;10% (pump) and &#60;15% (Stokes beams). If beams are not attenuated, the laser power can cause damage to the system and will burn biological samples.</li>
	<li>For cellular imaging, set the optimal laser settings to 6% (pump) and 10% (Stokes), corresponding to 12 mW and 30 mW at the sample.</li>
	<li>The SF-TRU has two settings: femtosecond (fs) mode and picosecond (ps) mode.</li>
	<li>For fs mode, push in the knobs on either side of the box (this removes the dispersion gratings from the beam path).</li>
	<li>For ps mode, pull out the knobs on either side of the box (this places the dispersion gratings in the beam path).</li>
	<li>For hyperspectral imaging, select the ps mode.</li>
	<li>Ensure that the delay stage is in the correct position for the pump and Stokes beams to be temporally overlapped for C-H imaging on this system.</li>
	<li>Ensure that the dispersion settings of the pump and Stokes beam are correct; for an 802 nm pump, this is 5 mm for the pump beam and 30 mm for the Stokes beam.</li>
</ol>
3. Modulating the Stokes beam for SRS imaging
<ol>
	<li>For SRS detection, modulate the intensity of the Stokes beam.</li>
	<li>Switch on the signal generator for the beam modulation.</li>
	<li>Recall previous settings, which are as follows: 
	OUTPUT 1: Square wave: Frequency = 19.5 MHz, Amplitude = 1.4 V. 
	OUTPUT 2: Square wave: Frequency = 19.5 MHz, Amplitude = 300 mV.</li>
	<li>Switch on Outputs 1 and 2.</li>
	<li>Connect Output 1 to an amplifier for the EOM in the SF-TRU box via a BNC cable.</li>
	<li>Connect Output 2 to the lock-in amplifier used for SRS detection via a BNC cable.</li>
	<li>Switch on the power supply to the amplifier on the gantry.</li>
</ol>
4. Switching on the lock-in amplifier
<ol>
	<li>For SRS imaging, use the SRS detection module that comprises a large area photodiode and a purpose-built lock-in amplifier.</li>
	<li>Switch on the power supply to the lock-in amplifier on the gantry.</li>
	<li>Load the software for the lock-in amplifier &#8220;SRS detection Module&#8221; on the PC.</li>
	<li>In the software, chose the following settings: Phase = 0&#176;, Offset = -80 mW, Gain = 58 dB.</li>
	<li>Select the lock-in amplifier integration time constant in the software: for cell imaging the time constant of 2 &#181;s gives good quality images.</li>
</ol>
5. Operating on the laser scanning microscope
<ol>
	<li>Switch on plugs of all the equipment except the remote-control box. Wait until standby is lit on the control box located on the top of the gantry.</li>
	<li>Switch on the remote-control box on the gantry and switch on the back of the box. Wait until the remote light goes blue (on the control box), and press Start Operation.</li>
	<li>Turn on the PC of the laser scanning microscope.</li>
	<li>Run the confocal microscope software.</li>
	<li>Check the light path of the microscope via the computer software. 
	NOTE: Some modifications have been made to the microscope to make it suitable for SRS imaging: (i) In the beam path, a 775 nm short pass has been added to the filter wheel where the laser beams enter the scan unit, this can be selected in the Lightpath tab in the computer software. (ii) Instead of using the confocal microscope&#8217;s inbuilt PMTs for detection, a bespoke detector has been added to the transmitted light path, which allows both SRS and CARS detection. (iii) The outputs from these detectors are connected to the Analog box on the gantry. CARS signal from the PMT is connected via a BNC cable to CD1 and SRS signal from the lock-in amplifier is connected via a BNC cable to CD2.</li>
	<li>Control the focus through the touch screen or the remote-control knobs or the computer software.</li>
	<li>Select the desired image settings in the software; this includes zoom, the image size in pixels and pixel dwell time.</li>
	<li>Ensure that the imaging speed (pixel dwell time) is greater than the integration time set in the lock-in amplifier software. 
	NOTE: A NA 1.2 water immersion objective was used for imaging.</li>
</ol>
6. Mounting a sample in the microscope stage
<ol>
	<li>Prepare the cell samples for imaging as follows.
	<ol>
		<li>Culture 4T1 mouse mammary carcinoma cells in RPMI-1640 supplemented with 10% Fetal Bovine Serum (FBS). Change the medium every 2 days and passage the cells at 70%&#8211;80% confluence. 
		NOTE: Passages 8&#8211;10 were used throughout the course of the experiments described here.</li>
		<li>For imaging experiments, incubate 1 x 105 cells on square coverslips for 24 h at 37&#8201;&#176;C, 5% CO2. Then, wash the cells with pre-warmed phosphate-buffered saline (PBS) and incubate with gold nanoparticles (1:100 dilution in cell culture medium, stock concentration: 2.3 x 1010 particles/mL, diameter: 60 nm) for 4 h.</li>
		<li>Prior to imaging, wash the cells with ice-cold PBS and fix them with 4% paraformaldehyde in PBS for 15 min at room temperature (RT). Wash the cells with PBS 3x, and then mount between two coverslips for imaging.</li>
	</ol>
	</li>
	<li>Place a droplet of distilled water on top of the water immersion objective that focuses the laser beams into to the sample between the objective and the sample. Then, place the sample on the microscope stage.</li>
	<li>A condenser with NA 1.4 collects the forward propagated light. Apply a droplet of water between the condenser and the sample for imaging. Adjust the height of the condenser to approximately 1 mm above the sample to collect the maximum amount of light.</li>
	<li>The SRS detector is on a moveable mount, which allows it to be slide in and out from the collection optical pathway. In order to focus on the sample using white light, move the SRS detector out of the beam path.</li>
	<li>Place the detector back in the beam path, ready for SRS imaging.</li>
</ol>
7. Changing the Raman shift and collecting a hyperspectral data stack
NOTE: The Raman shift at which the imaging takes place is dependent on the delay stage position in the SF-TRU box and the wavelength of the pump beam from the laser system.
<ol>
	<li>For large changes in the Raman shift, change the laser wavelength. For example, for imaging CH vibrations between 2,800&#8211;3,100 cm-1, select 802 nm, whereas, for imaging around 1,600 cm-1 for the amide I peak, select the pump beam at 898 nm. Adjust this via the software.</li>
	<li>To achieve small adjustments in the Raman shift, scan the delay stage in the SF-TRU unit.</li>
	<li>The SF-TRU gives delay stage positions in millimeters (mm). To convert the delay stage position from mm to cm-1, perform the calibration hyperspectral scan using a sample of polystyrene beads and compare the peak positions found in the calibration scan with those taken on a spontaneous Raman setup. This will provide a linear equation, which converts the delay stage position in mm to Raman shift in cm-1.</li>
	<li>To generate a hyperspectral scan, take a time series of images at different delay stage positions.</li>
	<li>To synchronize Image acquisition and movement of the delay stage, use the analog unit to send and receive TTL signals to and from the SF-TRU system. 
	NOTE: To do this, the analog box has the following connections: (i) TRIG OUT VD1&#8212;this is connected to the SF-TRU via a BNC and sends a TTL signal to tell the delay stage to move +1 increment. (ii) TRIG In 1&#8212;here, a signal is received via BNC when the delay stage has finished its movement, and this is used to trigger the next image in the time series.</li>
	<li>For the hyperspectral data set, select the following settings in the confocal microscope software:
	<ol>
		<li>In the Trigger tab, select Start by TTL trigger (ON) and Trigger (every frame).</li>
		<li>In the Series tab, set the time series ON and the z series OFF.</li>
		<li>Set the number of frames in the time series to match the number of steps in the ATM software (101 steps is used here).</li>
	</ol>
	</li>
	<li>In the ATM software, go to the Run tab.
	<ol>
		<li>Set the delay stage positions in millimeters for the start and stop of the hyperspectral scan. For CH high wavenumber region, use the start position = 90.25 mm and the stop position = 92.25 mm.</li>
		<li>Ensure that the number of steps matches the number of frames in the confocal microscope software.</li>
	</ol>
	</li>
	<li>After checking the correct settings, start the hyperspectral stack in the confocal microscope software first. Go to Acquire and click on Start Scan. Then, in the ATM software, click on Start. This initiates the collection of a series of images, with incremental increases in the delay stage position between the selected start and stop positions</li>
	<li>Export the hyperspectral image series as a multi-frame .tif file and use a suitable software package to carry out the calibration.</li>
	<li>Once the calibration is completed, use the linear calibration equation to convert the delay stage position to the Raman shifts.</li>
	<li>For switching between imaging in the CH vibrational region (2,800&#8211;3,100 cm-1) to the amide I vibrational region 1,500&#8211;1,700 cm-1), do the following.
	<ol>
		<li>Change the pump wavelength in the laser software from 802 nm to 898 nm.</li>
		<li>Change the Stokes dispersion setting in the ATM software from 30 mm to 5 mm.</li>
		<li>Make a small adjustment to the mirror in the pump beam dispersion pathway inside the SF-TRU box. This is done by adjusting the lower knob on the mirror mount, which changes the mirror angle by an incremental amount.</li>
		<li>When performing a hyperspectral scan over the amide I region, set the start and stop positions in the ATM software to 89 and 91mm, respectively 
		CAUTION: Laser safety goggles must be worn for this step as the laser beam will be exposed.</li>
	</ol>
	</li>
</ol>

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The authors declare no conflicts of interest.

This study has presented the combination of SF-TRU module and ultrafast dual-output laser system demonstrated its applications for multimodal microspectroscopy. With its ability to investigate gold nanoparticles&#8217; (AuNPs&#8217;) uptake by cancer cells, the multimodal imaging platform can visualize the cellular responses to hyperthermic cancer treatments when laser beams are absorbed by AuNPs.
Moreover, rapid chemically specific imaging and high spectral resolution are achieved by employin...

Gold nanoparticles (AuNPs) have shown great potential as biocompatible imaging probes, for example, as effective surface-enhanced Raman spectroscopy (SERS) substrates in various biomedical applications. Major applications include fields such as biosensing, bioimaging, surface-enhanced spectroscopies, and photothermal therapy for cancer treatment1. Furthermore, probing AuNPs in living systems is crucial to assessing and understanding the interaction between AuNPs and biological systems. There are various analytical techniques, including Fourier transform infrared (FTIR) spectroscopy2, laser ablation inductively coupled mass spectrometry (LA-ICP-MS)3, and magnetic resonance imaging (MRI)4 that have been successfully used to investigate the distribution of AuNPs in tissues. Nevertheless, these methods suffer from several drawbacks such as being time-consuming and involving complex sample preparation3, requiring long acquisition times, or the lack of sub-micron spatial resolution2,4.
Compared to conventional imaging techniques, nonlinear optical microscopy offers several advantages for probing live cells and AuNPs: The nonlinear optical microscopy achieves deeper imaging depth and provides intrinsic 3D optical sectioning capability with the use of near-IR ultrafast lasers. With the significant improvement of imaging speed and detection sensitivity, two-photon excited fluorescence (TPEF)5,6,7 and second harmonic generation (SHG)8,9,10 microscopy have been demonstrated to further improve non-invasive imaging of endogenous biomolecules in living cells and tissues. Moreover, utilizing novel pump-probe nonlinear optical techniques such as transient absorption (TA)11,12,13,14 and stimulated Raman scattering (SRS)15,16,17,18, it is possible to derive label-free biochemical contrast of cellular structures and AuNPs. Visualizing AuNPs without the use of extrinsic labels is of great importance since chemical perturbations of the nanoparticles will modify their physical properties and hence their uptake in cells.
This protocol presents the implementation of a Spectral Focusing Timing and Recombination Unit (SF-TRU) module for a dual-wavelength pulse laser, enabling fast multimodal imaging of AuNPs and cancer cells. This work aims to demonstrate the details of integrated TPEF, TA, and SRS techniques on a laser scanning microscope.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>APE SRS Detection Unit</td>
			<td>APE (Angewandte Physik &#38; Elektronik GmbH)</td>
			<td>APE Lock-in Module</td>
			<td>Combined system containing a large area Si photo-diode for detecting the pump beam along with a Lock-In amplifier for detecting the beam modulations</td>
		</tr>
		<tr>
			<td>Confocal Scanning Unit</td>
			<td>Olympus</td>
			<td>FV 3000</td>
			<td>Confocal scanning unit used for imaging</td>
		</tr>
		<tr>
			<td>CML Latex Beads, 4% w/v, 1.0 &#181;m</td>
			<td>Invitrogen</td>
			<td>C37483</td>
			<td>Polystyrene microspheres</td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>Thorlabs</td>
			<td>CG15CH2</td>
			<td>22 mm x 22 mm coverslips for seeding cells</td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Gibco</td>
			<td>10500-064</td>
			<td>Foetal Bovine Serum (Heat Inactivated)</td>
		</tr>
		<tr>
			<td>Flouview</td>
			<td>Olympus</td>
			<td>FV31S-SW</td>
			<td>Laser scanning microscope control software</td>
		</tr>
		<tr>
			<td>Function Generator</td>
			<td>BX precision</td>
			<td>40543</td>
			<td>Used to generate square wave function which is fed to EOM in SF-TRU to produce modulations in the stokes beam</td>
		</tr>
		<tr>
			<td>FV3000</td>
			<td>Olympus</td>
			<td>IX83P2ZF</td>
			<td>Other microscope frames can be used.</td>
		</tr>
		<tr>
			<td>Gold Nanoparticles</td>
			<td>Nanopartz</td>
			<td>A11-60</td>
			<td>Spherical gold nanoparticles, 60 nm diameter</td>
		</tr>
		<tr>
			<td>Input Output Interface</td>
			<td>Olympus</td>
			<td>FV30 ANALOG</td>
			<td>This unit allows voltage readouts from PMT and LockIn to be fed into the confocal scanning software and allows timing pulses to be sent between the olympus microscope and the SF-TRU unit.</td>
		</tr>
		<tr>
			<td>InSight X3</td>
			<td>Newport</td>
			<td>Spectra-Physics</td>
			<td>Dual-output femtosecond pulsed laser. Tunable (680&#8211;1300 nm) and fixed (1045 nm) laser outputs with the repetition rate of 80 MHz.</td>
		</tr>
		<tr>
			<td>Microscope Frame</td>
			<td>Olympus</td>
			<td>IX83</td>
			<td>Inverted microscope</td>
		</tr>
		<tr>
			<td>Mouse 4T1 cells</td>
			<td>ATCC</td>
			<td>CRL-2539</td>
			<td>Mouse breast cancer cells</td>
		</tr>
		<tr>
			<td>NA 1.2 Water Immersion Objective</td>
			<td>Olympus</td>
			<td>UPLSAPO60XW/IR</td>
			<td>The multiphoton 60x Objective has a 0.28 mm working distance. Other similar objectives can be used.</td>
		</tr>
		<tr>
			<td>NA 1.4 Condenser</td>
			<td>Nikon</td>
			<td>CSC1003</td>
			<td>Other condensers with NA higher than the excitation objective can also be used.</td>
		</tr>
		<tr>
			<td>PMT</td>
			<td>Hamamatsu</td>
			<td>R3896</td>
			<td>PMT used for detecting anti-stokes photos for CARS micrsocopy</td>
		</tr>
		<tr>
			<td>PMT Connector</td>
			<td>Hamamatsu</td>
			<td>C13654-01-Y002</td>
			<td>Connector for PMT</td>
		</tr>
		<tr>
			<td>Power Supply</td>
			<td>RS</td>
			<td>RSPD-3303 C</td>
			<td>Programmable power supply which is used for providing the correct voltage to the PMT</td>
		</tr>
		<tr>
			<td>RPMI-1640</td>
			<td>Gibco</td>
			<td>A10491-01</td>
			<td>Roswell Park Memorial Institute (RPMI) 1640 Medium has since been found suitable for a variety of mammalian cells.</td>
		</tr>
		<tr>
			<td>SF-TRU</td>
			<td>Newport Spectra Physics</td>
			<td>SF-TRU</td>
			<td>System designed for controlling the time delay and dispersion of the 2 laser outputs and for performing the beam modulations required for SRS</td>
		</tr>
	</tbody>
</table>

biomolecular imaging of cellular uptake of nanoparticles using multimodal nonlinear optical microscopy

Probing gold nanoparticles (AuNPs) in living systems is essential to reveal the interaction between AuNPs and biological tissues. Moreover, by integrating nonlinear optical signals such as stimulated Raman scattering (SRS), two-photon excited fluorescence (TPEF), and transient absorption (TA) into an imaging platform, it can be used to reveal biomolecular contrast of cellular structures and AuNPs in a multimodal manner. This article presents a multimodal nonlinear optical microscopy and applies it to perform chemically specific imaging of AuNPs in cancer cells. This imaging platform provides a novel approach for developing more efficient functionalized AuNPs and determining whether they are within vasculatures surrounding the tumor, pericellular, or cellular spaces.

The Spectral Focusing Timing and Recombination Unit (SF-TRU) module is introduced between the dual-output femtosecond laser and the modified laser scanning microscope. The tunable ultrafast laser system used in this study has two output ports delivering one beam at a fixed 1,045 nm wavelength and the other beam tunable in the range of 680&#8211;1,300 nm. A detailed schematic of the SF-TRU module and multimodal imaging platform is depicted in Figure 1. The SF-TRU is employed to chirp two femt...

Watch this Scientific Journal Video about Biomolecular Imaging of Cellular Uptake of Nanoparticles using Multimodal Nonlinear Optical Microscopy at JoVE.com

Biomolecular Imaging of Cellular Uptake of Nanoparticles using Multimodal Nonlinear Optical Microscopy

This article presents the integration of a spectral-focusing module and a dual-output pulse laser, enabling rapid hyperspectral imaging of gold nanoparticles and cancer cells. This work aims to demonstrate the details of multimodal nonlinear optical techniques on a standard laser scanning microscope.

Probing gold nanoparticles (AuNPs) in living systems is essential to reveal the interaction between AuNPs and biological tissues. Moreover, by integrating ...

biomolecular-imaging-cellular-uptake-nanoparticles-using-multimodal

Research

JoVE Journal

Bioengineering

1.8K Views.  University of Exeter. This article presents the integration of a spectral-focusing module and a dual-output pulse laser, enabling rapid hyperspectral imaging of gold nanoparticles and cancer cells. This work aims to demonstrate the details of multimodal nonlinear optical techniques on a standard laser scanning microscope.

Video: Biomolecular Imaging of Cellular Uptake of Nanoparticles using Multimodal Nonlinear Optical Microscopy

Biomolecular Detection employing the Interferometric Reflectance Imaging Sensor (IRIS)

The sensitive measurement of biomolecular interactions has use in many fields and industries such as basic biology and microbiology, environmental/agricultural/biodefense monitoring, nanobiotechnology, and more. For diagnostic applications, monitoring (detecting) the presence, absence, or abnormal expression of targeted proteomic or genomic biomarkers found in patient samples can be used to determine treatment approaches or therapy efficacy. In the research arena, information on molecular affinities and specificities are useful for fully characterizing the systems under investigation.
Many of the current systems employed to determine molecular concentrations or affinities rely on the use of labels. Examples of these systems include immunoassays such as the enzyme-linked immunosorbent assay (ELISA), polymerase chain reaction (PCR) techniques, gel electrophoresis assays, and mass spectrometry (MS). Generally, these labels are fluorescent, radiological, or colorimetric in nature and are directly or indirectly attached to the molecular target of interest. Though the use of labels is widely accepted and has some benefits, there are drawbacks which are stimulating the development of new label-free methods for measuring these interactions. These drawbacks include practical facets such as increased assay cost, reagent lifespan and usability, storage and safety concerns, wasted time and effort in labelling, and variability among the different reagents due to the labelling processes or labels themselves. On a scientific research basis, the use of these labels can also introduce difficulties such as concerns with effects on protein functionality/structure due to the presence of the attached labels and the inability to directly measure the interactions in real time.
Presented here is the use of a new label-free optical biosensor that is amenable to microarray studies, termed the Interferometric Reflectance Imaging Sensor (IRIS), for detecting proteins, DNA, antigenic material, whole pathogens (virions) and other biological material. The IRIS system has been demonstrated to have high sensitivity, precision, and reproducibility for different biomolecular interactions [1-3]. Benefits include multiplex imaging capacity, real time and endpoint measurement capabilities, and other high-throughput attributes such as reduced reagent consumption and a reduction in assay times. Additionally, the IRIS platform is simple to use, requires inexpensive equipment, and utilizes silicon-based solid phase assay components making it compatible with many contemporary surface chemistry approaches.
Here, we present the use of the IRIS system from preparation of probe arrays to incubation and measurement of target binding to analysis of the results in an endpoint format. The model system will be the capture of target antibodies which are specific for human serum albumin (HSA) on HSA-spotted substrates.

Biomolecular Detection employing the Interferometric Reflectance Imaging Sensor IRIS

Quantitative, high-throughput, real-time, and label-free biomolecular detection (DNA, protein, etc.) on SiO2 surfaces can be achieved using a simple interferometric technique which relies on LED illumination, minimal optical components, and a camera. The Interferometric Reflectance Imaging Sensor (IRIS) is inexpensive, simple to use, and amenable to microarray formats.

The sensitive measurement of biomolecular interactions has use in many fields and industries such as basic biology and microbiology, ...

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Analyzing Cellular Internalization of Nanoparticles and Bacteria by Multi-spectral Imaging Flow Cytometry

Nanoparticulate systems have emerged as valuable tools in vaccine delivery through their ability to efficiently deliver cargo, including proteins, to antigen presenting cells1-5. Internalization of nanoparticles (NP) by antigen presenting cells is a critical step in generating an effective immune response to the encapsulated antigen. To determine how changes in nanoparticle formulation impact function, we sought to develop a high throughput, quantitative experimental protocol that was compatible with detecting internalized nanoparticles as well as bacteria. To date, two independent techniques, microscopy and flow cytometry, have been the methods used to study the phagocytosis of nanoparticles. The high throughput nature of flow cytometry generates robust statistical data. However, due to low resolution, it fails to accurately quantify internalized versus cell bound nanoparticles. Microscopy generates images with high spatial resolution; however, it is time consuming and involves small sample sizes6-8. Multi-spectral imaging flow cytometry (MIFC) is a new technology that incorporates aspects of both microscopy and flow cytometry that performs multi-color spectral fluorescence and bright field imaging simultaneously through a laminar core. This capability provides an accurate analysis of fluorescent signal intensities and spatial relationships between different structures and cellular features at high speed.
	Herein, we describe a method utilizing MIFC to characterize the cell populations that have internalized polyanhydride nanoparticles or Salmonella enterica serovar Typhimurium. We also describe the preparation of nanoparticle suspensions, cell labeling, acquisition on an ImageStreamX system and analysis of the data using the IDEAS application. We also demonstrate the application of a technique that can be used to differentiate the internalization pathways for nanoparticles and bacteria by using cytochalasin-D as an inhibitor of actin-mediated phagocytosis.

In this article, we describe a method utilizing multi-spectral imaging flow cytometry to quantify the internalization of polyanhydride nanoparticles or bacteria by RAW 264.7 cells.

Nanoparticulate  systems have emerged as valuable tools in vaccine delivery through their  ability to efficiently deliver cargo, including proteins, to ...

Quantitative Optical Microscopy: Measurement of Cellular Biophysical Features with a Standard Optical Microscope

We describe the use of a standard optical microscope to perform quantitative measurements of mass, volume, and density on cellular specimens through a combination of bright field and differential interference contrast imagery. Two primary approaches are presented: noninterferometric quantitative phase microscopy (NIQPM), to perform measurements of total cell mass and subcellular density distribution, and Hilbert transform differential interference contrast microscopy (HTDIC)&#160;to determine volume. NIQPM is based on a simplified model of wave propagation, termed the paraxial approximation, with three underlying assumptions: low numerical aperture (NA) illumination, weak scattering, and weak absorption of light by the specimen. Fortunately, unstained cellular specimens satisfy these assumptions and low NA illumination is easily achieved on commercial microscopes. HTDIC is used to obtain volumetric information from through-focus DIC imagery under high NA illumination conditions. High NA illumination enables enhanced sectioning of the specimen along the optical axis. Hilbert transform processing on the DIC image stacks greatly enhances edge detection algorithms for localization of the specimen borders in three dimensions by separating the gray values of the specimen intensity from those of the background. The primary advantages of NIQPM and HTDIC lay in their technological accessibility using &#8220;off-the-shelf&#8221; microscopes. There are two basic limitations of these methods: slow z-stack acquisition time on commercial scopes currently abrogates the investigation of phenomena faster than 1 frame/minute, and secondly, diffraction effects restrict the utility of NIQPM and HTDIC to objects from 0.2 up to 10 (NIQPM) and 20 (HTDIC) &#956;m in diameter, respectively. Hence, the specimen and its associated time dynamics of interest must meet certain size and temporal constraints to enable the use of these methods. Excitingly, most fixed cellular specimens are readily investigated with these methods.

We describe the use of a standard optical microscope to perform quantitative measurements of cellular mass, volume, and density through a combination of bright field and differential interference contrast imagery.

We describe the use of a standard optical microscope to perform quantitative measurements of mass, volume, and density on cellular specimens through a ...

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous number of imaging techniques currently available and the progress in instrumentation, there is still a need for highly sensitive probes that are suitable for in vivo imaging. One typical problem of available preclinical fluorescent probes is their rapid clearance in vivo, which reduces their imaging sensitivity. To circumvent rapid clearance, increase number of dye molecules at the target site, and thereby reduce background autofluorescence, encapsulation of the near-infrared fluorescent dye, DY-676-COOH in liposomes and verification of its potential for in vivo imaging of inflammation was done. DY-676 is known for its ability to self-quench at high concentrations. We first determined the concentration suitable for self-quenching, and then encapsulated this quenching concentration into the aqueous interior of PEGylated liposomes. To substantiate the quenching and activation potential of the liposomes we use a harsh freezing method which leads to damage of liposomal membranes without affecting the encapsulated dye. The liposomes characterized by a high level of fluorescence quenching were termed Lip-Q. We show by experiments with different cell lines that uptake of Lip-Q is predominantly by phagocytosis which in turn enabled the characterization of its potential as a tool for in vivo imaging of inflammation in mice models. Furthermore, we use a zymosan-induced edema model in mice to substantiate the potential of Lip-Q in optical imaging of inflammation in vivo. Considering possible uptake due to inflammation-induced enhanced permeability and retention (EPR) effect, an always-on liposome formulation with low, non-quenched concentration of DY-676-COOH (termed Lip-dQ) and the free DY-676-COOH were compared with Lip-Q in animal trials.

Fluorescence-quenching of a Liposomal-encapsulated Near-infrared Fluorophore as a Tool for In Vivo Optical Imaging

The use of fluorophores for in vivo imaging can be greatly limited by opsonization, rapid clearance, low detection sensitivity and cytotoxic effects on the host. Encapsulation of fluorophores in liposomes by film hydration and extrusion leads to fluorescence quenching and protection which enables in vivo imaging with high detection sensitivity.

Optical imaging offers a wide range of diagnostic modalities and has attracted a lot of interest as a tool for biomedical imaging. Despite the enormous ...

Biofunctionalized Prussian Blue Nanoparticles for Multimodal Molecular Imaging Applications

Multimodal, molecular imaging allows the visualization of biological processes at cellular, subcellular, and molecular-level resolutions using multiple, complementary imaging techniques. These imaging agents facilitate the real-time assessment of pathways and mechanisms in vivo, which enhance both diagnostic and therapeutic efficacy. This article presents the protocol for the synthesis of biofunctionalized Prussian blue nanoparticles (PB NPs) - a novel class of agents for use in multimodal, molecular imaging applications. The imaging modalities incorporated in the nanoparticles, fluorescence imaging and magnetic resonance imaging (MRI), have complementary features. The PB NPs possess a core-shell design where gadolinium and manganese ions incorporated within the interstitial spaces of the PB lattice generate MRI contrast, both in T1 and T2-weighted sequences. The PB NPs are coated with fluorescent avidin using electrostatic self-assembly, which enables fluorescence imaging. The avidin-coated nanoparticles are modified with biotinylated ligands that confer molecular targeting capabilities to the nanoparticles. The stability and toxicity of the nanoparticles are measured, as well as their MRI relaxivities. The multimodal, molecular imaging capabilities of these biofunctionalized PB NPs are then demonstrated by using them for fluorescence imaging and molecular MRI in vitro.

This protocol describes the synthesis of biofunctionalized Prussian blue nanoparticles and their use as multimodal, molecular imaging agents. The nanoparticles have a core-shell design where gadolinium or manganese ions within the nanoparticle core generate MRI contrast. The biofunctional shell contains fluorophores for fluorescence imaging and targeting ligands for molecular targeting.

Multimodal, molecular imaging allows the visualization of biological processes at cellular, subcellular, and molecular-level resolutions using multiple, ...

Correlative Light- and Electron Microscopy Using Quantum Dot Nanoparticles

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using widefield fluorescence light microscopy and transmission electron microscopy (TEM). To demonstrate the protocol we have immunolabeled ultrathin epoxy sections of human somatostatinoma tumor using a primary antibody to somatostatin, followed by a biotinylated secondary antibody and visualization with streptavidin conjugated 585 nm cadmium-selenium (CdSe) quantum dots (QDs). The sections are mounted on a TEM specimen grid then placed on a glass slide for observation by widefield fluorescence light microscopy. Light microscopy reveals 585 nm QD labeling as bright orange fluorescence forming a granular pattern within the tumor cell cytoplasm. At low to mid-range magnification by light microscopy the labeling pattern can be easily recognized and the level of non-specific or background labeling assessed. This is a critical step for subsequent interpretation of the immunolabeling pattern by TEM and evaluation of the morphological context. The same section is then blotted dry and viewed by TEM. QD probes are seen to be attached to amorphous material contained in individual secretory granules. Images are acquired from the same region of interest (ROI) seen by light microscopy for correlative analysis. Corresponding images from each modality may then be blended to overlay fluorescence data on TEM ultrastructure of the corresponding region.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of epoxy embedded human pathology tissue. We employ commercial antibody fragment conjugated QDs that are visualized by widefield fluorescence light microscopy and transmission electron microscopy.

A method is described whereby quantum dot (QD) nanoparticles can be used for correlative immunocytochemical studies of human pathology tissue using ...

Targeted Plasma Membrane Delivery of a Hydrophobic Cargo Encapsulated in a Liquid Crystal Nanoparticle Carrier

The controlled delivery of drug/imaging agents to cells is critical for the development of therapeutics and for the study of cellular signaling processes. Recently, nanoparticles (NPs) have shown significant promise in the development of such delivery systems. Here, a liquid crystal NP (LCNP)-based delivery system has been employed for the controlled delivery of a water-insoluble dye, 3,3&#8242;-dioctadecyloxacarbocyanine perchlorate (DiO), from within the NP core to the hydrophobic region of a plasma membrane bilayer. During the synthesis of the NPs, the dye was efficiently incorporated into the hydrophobic LCNP core, as confirmed by multiple spectroscopic analyses. Conjugation of a PEGylated cholesterol derivative to the NP surface (DiO-LCNP-PEG-Chol) enabled the binding of the dye-loaded NPs to the plasma membrane in HEK 293T/17 cells. Time-resolved laser scanning confocal microscopy and F&#246;rster resonance energy transfer (FRET) imaging confirmed the passive efflux of DiO from the LCNP core and its insertion into the plasma membrane bilayer. Finally, the delivery of DiO as a LCNP-PEG-Chol attenuated the cytotoxicity of DiO; the NP form of DiO exhibited ~30-40% less toxicity compared to DiOfree delivered from bulk solution. This approach demonstrates the utility of the LCNP platform as an efficient modality for the membrane-specific delivery and modulation of hydrophobic molecular cargos.

A liquid crystal nanoparticle (LCNP) nanocarrier is exploited as a vehicle for the controlled delivery of a hydrophobic cargo to the plasma membrane of living cells.

The controlled delivery of drug/imaging agents to cells is critical for the development of therapeutics and for the study of cellular signaling processes. ...

Long-term Live Imaging Device for Improved Experimental Manipulation of Zebrafish Larvae

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for live high-resolution microscopic imaging of dynamic biological phenomena in space and time with cellular resolution. However, the traditional method of agarose encapsulation for live imaging can impede larval development and tissue regrowth. Therefore, this manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which was designed and fabricated as a functionally compartmentalized device to orient larvae for high-resolution microscopy while permitting caudal fin transection within the device and subsequent unrestrained tail development and re-growth. This device allows for wounding and long-term imaging while maintaining viability. Given that the zWEDGI mold is 3D printed, the customizability of its geometries make it easily modified for diverse zebrafish imaging applications. Furthermore, the zWEDGI offers numerous benefits, such as access to the larva during experimentation for wounding or for the application of reagents, paralleled orientation of multiple larvae for streamlined imaging, and reusability of the device.

This manuscript describes the zWEDGI (zebrafish Wounding and Entrapment Device for Growth and Imaging), which is a compartmentalized device designed to orient and restrain zebrafish larvae. The design permits tail transection and long-term collection of high-resolution fluorescent microscopy images of wound healing and regeneration.

The zebrafish larva is an important model organism for both developmental biology and wound healing. Further, the zebrafish larva is a valuable system for ...