The authors appreciate Dr. Lynnea Brumbaugh's close reading of the manuscript. This work was sponsored by National Institutes of Health Grants R01 EB000712, R01 EB008085, R01 CA134539, U54 CA136398, and 5P60 DK02057933. Prof. Lihong V. Wang has a financial interest in Microphotoacoustics, Inc. and Endra, Inc., which, however, did not support this work.

1. System configuration
<ol>
<li>Optical irradiation
<ol>
 <li>Optical irradiation source: a diode-pumped solid-state pulsed laser (INNOSLAB, Edgewave) and a dye laser (CBR-D, Sirah). </li>
 <li> The output laser beam (pulse width: 7 ns) is focused by a condenser lens (LA1131, Thorlabs) to pass through a 50-&mu;m pinhole (P50C, Thorlabs).</li>
 <li>The pinhole is positioned slightly away from the focus of the condenser lens to match the pinhole diameter with the fundamental-mode beam diameter for effective spatial filtering. </li>
 <li>The filtered beam is attenuated by a neutral-density filter (NDC-50C-2M, Thorlabs) and then coupled into a single-mode optical fiber (P1-460A-FC-2, Thorlabs). </li>
 <li>The fiber output fills the back aperture of a microscope objective (RMS4X, Thorlabs) to achieve a diffraction-limited optical focus of ~2.6 &mu;m at the wavelength of 570 nm. </li> 
</ol>
</li>
<li>Ultrasonic detection
<ol>
<li>Ultrasonic transducer: 50-MHz central frequency (V214-BB-RM, Olympus-NDT). </li>
<li>The ultrasonic transducer is attached to a home-made acoustic-optical beam combiner11 for ultrasonic detection, which is aligned coaxially with the diffraction-limited optical irradiation. </li>
<li>A spherical cavity is ground out of the bottom of the combiner to produce an acoustic lens. This acoustic lens has a numerical aperture of 0.5 in water and gives an acoustic focal diameter of 43 &mu;m at the 50-MHz central frequency. </li>
<li>The optical and acoustic foci are aligned confocally to maximize the detection sensitivity.</li>
</ol>
</li>
<li>Acoustic coupling
<ol>
 <li>Dry ultrasonic coupling is employed to avoid submerging experimental animals in water, which was used in early photoacoustic imaging systems12. </li>
 <li>An imaging window is opened in the bottom of a Petri dish (9 cm in diameter) and is sealed with an ultrasonically and optically transparent polyethylene membrane. </li>
 <li>Ultrasonic gel (Clear Image, SonoTech) between the polyethylene membrane and the object to be imaged couples the generated photoacoustic wave from the object to the Petri dish, and deionized water in the Petri dish further couples the wave to the submerged OR-PAM imaging head. </li> 
</ol>
</li>
<li>Electronics
<ol>
 <li>The photoacoustic signal detected by the ultrasonic transducer is amplified by two cascaded amplifiers (ZFL 500LN, Mini-Circuits) </li>
 <li>The amplified signal is digitized by a 14-bit data acquisition (DAQ) board (CompuScope 14200, Gage Applied Sciences) at a sampling rate of 200 MS/s. </li>
</ol>
</li>
<li>Scanning scheme
<ol>
 <li>Two-dimensional (2-D) raster scanning of the OR-PAM imaging head along the horizontal (x-y) plane is controlled by a personal computer, which triggers both the DAQ board and the pump laser. The trigger signal is synchronized with the clock-out signal from the DAQ board. </li>
 <li>The fast axis of the 2-D scanner is defined as the direction of the cross-sectional scan (B-scan).</li>
 <li>A sequence of B-scan images can be acquired by translating the imaging head along the slow axis to form a volumetric image, which can be viewed either in a direct 3-D rendering or in a 2-D maximum amplitude projection (MAP) image.</li> 
</ol>
</li>
</ol>
2. System alignment
<ol>
 <li>Use pulse-echo ultrasound and an ultrasonic reflector to determine the position of the acoustic focal plane (i.e., the time delay from the trigger signal to the maximum pulse-echo ultrasonic signal). This step is required to be performed only once when building the OR-PAM system. </li>
 <li>Maximize the coupling efficiency of the single-mode fiber.</li>
 <li>Apply ultrasonic gel on top of an optically absorbing object (e.g., a piece of black tape) and gently attach it beneath the imaging window in the Petri dish filled with deionized water. </li>
 <li>Lower the imaging head into the water, and remove air bubbles trapped under the acoustic lens. </li>
 <li>Adjust the imaging head until the photoacoustic signal of the absorbing object is from the acoustic focal plane, which can be judged from the acoustic delay. </li>
 <li>Adjust the vertical position (i.e., z position) of the microscope objective to maximize the amplitude of the photoacoustic signal generated from the flat object. The maximized signal amplitude suggests that the optical focus is aligned with the acoustic focus in the vertical direction. </li>
 <li>Adjust the horizontal positions (i.e., x and y positions) of the microscope objective until the photoacoustic signal generated from the target shows a symmetric pattern. The symmetry suggests that the optical focus is aligned with the acoustic focus in the horizontal direction.</li>
 <li>Repeat steps 2.6 and 2.7 until the photoacoustic signal is optimized in both symmetry and amplitude.</li>
</ol>
3. A sample experimental procedure-in vivo OR-PAM of mouse ear vasculature
<ol>
 <li>This step is not required for nude mice. Anesthetize the animal with an intraperitoneal injection of a cocktail [recipe: 1 ml ketamine (100 mg/ml), 0.1 ml xylazine (100 mg/ml), and 8.9 ml saline; dosage: 0.1 ml/10 g]. Shave the hair in the ear, and further depilate the residual hair with Surgi Cream (Category #: 82565, American International Industries) before cleaning it with deionized water. Note that such depilation might slightly irritate the skin vasculature and thus is best performed 24 hours before the planned experiment. </li>
 <li>Turn on the photoacoustic laser system, and check the system alignment. </li>
 <li>Anesthetize the mouse with 3% isoflurane vaporized by the inhalation gas (the typical flow rate is 1.0-1.5 l/min, depending on the animal's body weight), and 
 maintain the anesthesia with 1% isoflurane throughout the experiment. Medical-grade air is recommended as the inhalation gas to maintain the mouse at normal 
 physiological status. </li>
 <li>Transfer the mouse to a stereotactic stage, and control its body temperature at 37 &deg;C with a heating pad. </li>
 <li>Flatten the mouse ear on a plastic plate and apply a layer of ultrasound gel on top of the ear. Avoid trapping air bubbles inside the gel. 
 Then, place the ear under the imaging window and slowly raise the animal stage until the ultrasound gel contacts the bottom of the polyethylene membrane. 
 Soft contact is required because pressing the ear against the membrane may affect blood flow in the ear. </li>
 <li>Clamp a pulse oximeter to the mouse leg or tail to monitor its physiological status, and apply ointment to the eyes to prevent dryness and accidental laser damage to the mouse eyes.</li>
 <li>Lower the imaging head until the acoustic lens is immersed in deionized water, and remove air bubbles trapped under the acoustic lens. </li>
 <li>Check the laser fluence to make sure it is within the laser safety standards of the American National Standards Institute13. The laser fluence should not exceed 20 mJ/cm2, which translates into 80 nJ of laser pulse energy when the laser beam is focused at 150 &mu;m beneath the skin surface.</li>
 <li>Set the laser to the external-trigger mode and start trial scanning. Adjust the z position of the imaging head until the strongest photoacoustic signal is from the acoustic focal plane. </li>
 <li>Set correct scanning parameters and start formal image acquisition.</li>
 <li>After the experiment, turn off the laser, lift the imaging head out of the deionized water, lower the animal stage to release the mouse, clean the mouse ear with deionized water, turn off the anesthesia system and the temperature controller, and unload the mouse from the stereotactic stage. </li>
 <li>If repetitive imaging is required, put the mouse in an incubator with the environmental temperature set at 37 &deg;C. Return the mouse to the animal facility after it wakes up naturally. Otherwise, follow the animal protocols to euthanatize and dispose of it. </li> 
</ol>
4. Functional OR-PAM of total concentration and oxygen saturation of hemoglobin
<ol>
 <li>Oxyhemoglobin (HbO2) and deoxyhemoglobin (HbR) are the two major forms of hemoglobin, the predominant endogenous photoacoustic source in the visible spectral range. HbO2 and HbR have distinct optical absorption spectra and thus can be spectrally differentiated to quantify both the total concentration (HbT) and oxygen saturation (sO2) of hemoglobin5. Here are two guidelines to help select proper optical wavelengths for sO2 measurements: </li>
 <li>Wavelengths should be selected within the Q-band of the hemoglobin absorption spectrum (i.e., 550-600 nm) to ensure a sufficient signal-to-noise ratio and adequate penetration.</li>
 <li>Wavelengths where the absorption coefficients of HbO2 and HbR have a pronounced difference (e.g., HbR-dominant 561 nm and HbO2-dominant 578 nm) are recommended. </li> 
</ol>
Besides sO2, HbT can be calculated by adding [HbR] and [HbO2] together, or it can be directly measured at isosbestic wavelengths of the molar extinction coefficient spectra of hemoglobin (e.g., 530 nm, 545 nm, 570 nm, and 584 nm)14, where HbR and HbO2 have equal molar extinction coefficients.
5. Representative Results:
Shown in Figure 1 is the vascular anatomy and sO2 in a living nude mouse ear imaged by dual-wavelength (561 and 570 nm) OR-PAM. The typical image acquisition time for dual-wavelength sO2 measurements of such a region of interest (image size: 10 mm x 10 mm; step size: 2.5 &mu;m x 5 &mu;m) is ~80 min.
<img src="/files/ftp_upload/2729/2729fig1.jpg" alt="Figure 1" /> 
Figure 1. In vivo optical-resolution photoacoustic microscopy. MAP images of (A) the total hemoglobin concentration showing the vascular anatomy (acquired at 570 nm) and (B) the hemoglobin oxygen saturation (acquired at 561 nm and 570 nm) in a nude mouse ear. (C) Close-up of the boxed region in panel (A). The scale bar in panel (A) applies to both (A) and (B).

<ol>
	<li>Maslov, K., Zhang, H. F., Hu, S., Wang, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical-resolution+photoacoustic+microscopy+for+in+vivo+imaging+of+single+capillaries.">Optical-resolution photoacoustic microscopy for in vivo imaging of single capillaries.</a> Opt. Lett. 33, 929-931 (2008).</li><li>Zhang, C., Maslov, K., Wang, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subwavelength-resolution+label-free+photoacoustic+microscopy+of+optical+absorption+in+vivo.">Subwavelength-resolution label-free photoacoustic microscopy of optical absorption in vivo.</a> Opt. Lett. 35, 3195-3197 (2010).</li><li>Wang, Y., Maslov, K., Kim, C., Hu, S., Wang, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Integrated+photoacoustic+and+fluorescence+confocal+microscopy.">Integrated photoacoustic and fluorescence confocal microscopy.</a> IEEE. Trans. Biomed. Eng. 57, 2576-2578 (2010).</li><li>Jiao, S., Xie, Z., Zhang, H. F., Puliafito, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+multimodal+imaging+with+integrated+photoacoustic+microscopy+and+optical+coherence+tomography.">Simultaneous multimodal imaging with integrated photoacoustic microscopy and optical coherence tomography.</a> Opt. Lett. 34, 2961-2963 (2009).</li><li>Hu, S., Maslov, K., Tsytsarev, V., Wang, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+transcranial+brain+imaging+by+optical-resolution+photoacoustic+microscopy.">Functional transcranial brain imaging by optical-resolution photoacoustic microscopy.</a> J. Biomed. Opt. 14, 040503-040503 (2009).</li><li>Hu, S., Yan, P., Maslov, K., Lee, J. M., Wang, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intravital+imaging+of+amyloid+plaques+in+a+transgenic+mouse+model+using+optical-resolution+photoacoustic+microscopy.">Intravital imaging of amyloid plaques in a transgenic mouse model using optical-resolution photoacoustic microscopy.</a> Opt. Lett. 34, 3899-3901 (2009).</li><li>Hu, S., Rao, B., Maslov, K., Wang, L. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+Photoacoustic+Ophthalmic+Angiography.">Label-free Photoacoustic Ophthalmic Angiography.</a> Opt. Lett. 35, 1-3 (2010).</li><li>Jiao, S. L., Jiang, M. S., Hu, J. M., Fawzi, A., Zhou, Q. F., Shung, K. K., Puliafito, C. A., Zhang, H. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photoacoustic+ophthalmoscopy+for+in+vivo+retinal+imaging.">Photoacoustic ophthalmoscopy for in vivo retinal imaging.</a> Opt. Express. 18, 3967-3972 (2010).</li><li>Oladipupo, S., Hu, S., Santeford, A., Yao, J., Kovalski, J. R., Shohet, R., Maslov, K., Wang, L. V., Arbeit, J. 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All experimental animal procedures were carried out in conformance with the laboratory animal protocol approved by the School of Medicine Animal Studies Committee of Washington University in St. Louis.

In this video, we provide a detailed instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, and typical experimental procedures. Label-free, noninvasive OR-PAM has enabled studies of microvascular functioning and metabolism on a single capillary basis and thereby holds the potential to expand our understanding of microcirculation-related physiology and pathology. Microphotoacoustics is currently manufacturing this OR-PAM system.

Home-made acoustic-optical beam combiner:
<ul>
			<li>right-angle prism (NT32-545, Edmund Optics)</li>
			<li>rhomboid prism (NT49-419, Edmund Optics)</li>
			<li>silicone oil (1000cSt, Clearco Products)</li>
			<li>OR-PAM system (Microphotoacoustics)</li>
</ul>

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.

Watch this Scientific Journal Video about Three-dimensional Optical-resolution Photoacoustic Microscopy at JoVE.com

Three-dimensional Optical-resolution Photoacoustic Microscopy

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 ...

three-dimensional-optical-resolution-photoacoustic-microscopy

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17.7K Views.  Washington University in St. Louis. 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.

Video: Three-dimensional Optical-resolution Photoacoustic Microscopy

Alginate Hydrogels for Three-Dimensional Organ Culture of Ovaries and Oviducts

Ovarian cancer is the fifth leading cause of cancer deaths in women and has a 63% mortality rate in the United States1. The cell type of origin for ovarian cancers is still in question and might be either the ovarian surface epithelium (OSE) or the distal epithelium of the fallopian tube fimbriae2,3. Culturing the normal cells as a primary culture in vitro will enable scientists to model specific changes that might lead to ovarian cancer in the distinct epithelium, thereby definitively determining the cell type of origin. This will allow development of more accurate biomarkers, animal models with tissue-specific gene changes, and better prevention strategies targeted to this disease.
Maintaining normal cells in alginate hydrogels promotes short term in vitro culture of cells in their three-dimensional context and permits introduction of plasmid DNA, siRNA, and small molecules. By culturing organs in pieces that are derived from strategic cuts using a scalpel, several cultures from a single organ can be generated, increasing the number of experiments from a single animal. These cuts model aspects of ovulation leading to proliferation of the OSE, which is associated with ovarian cancer formation. Cell types such as the OSE that do not grow well on plastic surfaces can be cultured using this method and facilitate investigation into normal cellular processes or the earliest events in cancer formation4.
Alginate hydrogels can be used to support the growth of many types of tissues5. Alginate is a linear polysaccharide composed of repeating units of &beta;-D-mannuronic acid and &alpha;-L-guluronic acid that can be crosslinked with calcium ions, resulting in a gentle gelling action that does not damage tissues6,7. Like other three-dimensional cell culture matrices such as Matrigel, alginate provides mechanical support for tissues; however, proteins are not reactive with the alginate matrix, and therefore alginate functions as a synthetic extracellular matrix that does not initiate cell signaling5. The alginate hydrogel floats in standard cell culture medium and supports the architecture of the tissue growth in vitro.
A method is presented for the preparation, separation, and embedding of ovarian and oviductal organ pieces into alginate hydrogels, which can be maintained in culture for up to two weeks. The enzymatic release of cells for analysis of proteins and RNA samples from the organ culture is also described. Finally, the growth of primary cell types is possible without genetic immortalization from mice and permits investigators to use knockout and transgenic mice.

Culture of normal cells in their three-dimensional context represents an alternative method to study early events required for cellular transformation and tumorigenesis. This method is used to grow normal ovarian and oviductal cells to study early events in ovarian cancer formation.

Ovarian cancer is the fifth leading cause of cancer deaths in women and has a 63% mortality rate in the United States1. The cell type of origin for ...

Planar and Three-Dimensional Printing of Conductive Inks

Printed electronics rely on low-cost, large-area fabrication routes to create flexible or multidimensional electronic, optoelectronic, and biomedical devices1-3. In this paper, we focus on one- (1D), two- (2D), and three-dimensional (3D) printing of conductive metallic inks in the form of flexible, stretchable, and spanning microelectrodes.
Direct-write assembly4,5 is a 1-to-3D printing technique that enables the fabrication of features ranging from simple lines to complex structures by the deposition of concentrated inks through fine nozzles (~0.1 - 250 &mu;m). This printing method consists of a computer-controlled 3-axis translation stage, an ink reservoir and nozzle, and 10x telescopic lens for visualization. Unlike inkjet printing, a droplet-based process, direct-write assembly involves the extrusion of ink filaments either in- or out-of-plane. The printed filaments typically conform to the nozzle size. Hence, microscale features (&lt; 1 &mu;m) can be patterned and assembled into larger arrays and multidimensional architectures.
In this paper, we first synthesize a highly concentrated silver nanoparticle ink for planar and 3D printing via direct-write assembly. Next, a standard protocol for printing microelectrodes in multidimensional motifs is demonstrated. Finally, applications of printed microelectrodes for electrically small antennas, solar cells, and light-emitting diodes are highlighted.

Planar and three-dimensional printing of conductive metallic inks is described. Our approach provides new avenues for fabricating printed electronic, optoelectronic, and biomedical devices in unusual layouts at the microscale.

Printed electronics rely on low-cost, large-area fabrication routes to create flexible or multidimensional electronic, optoelectronic, and biomedical ...

Photoacoustic Cystography

Conventional pediatric cystography, which is based on diagnostic X-ray using a radio-opaque dye, suffers from the use of harmful ionizing radiation. The risk of bladder cancers in children due to radiation exposure is more significant than many other cancers. Here we demonstrate the feasibility of nonionizing and noninvasive photoacoustic (PA) imaging of urinary bladders, referred to as photoacoustic cystography (PAC), using near-infrared (NIR) optical absorbents (i.e. methylene blue, plasmonic gold nanostructures, or single walled carbon nanotubes) as an optical-turbid tracer. We have successfully imaged a rat bladder filled with the optical absorbing agents using a dark-field confocal PAC system. After transurethral injection of the contrast agents, the rat's bladders were photoacoustically visualized by achieving significant PA signal enhancement. The accumulation was validated by spectroscopic PA imaging. Further, by using only a laser pulse energy of less than 1 mJ/cm2 (1/20 of the safety limit), our current imaging system could map the methylene-blue-filled-rat-bladder at the depth of beyond 1 cm in biological tissues in vivo. Both in vivo and ex vivo PA imaging results validate that the contrast agents were naturally excreted via urination. Thus, there is no concern regarding long-term toxic agent accumulation, which will facilitate clinical translation.

Photoacoustic cystography (PAC) has a great potential to map urinary bladders, a radiation sensitive internal organ in pediatric patients, without using any ionizing radiation or toxic contrast agent. Here we demonstrate the use of PAC for mapping urinary bladders with an injection of optical-opaque tracers in rats in vivo.

Conventional pediatric cystography, which is based on diagnostic X-ray using a radio-opaque dye, suffers from the use of harmful ionizing radiation. The ...

Human Cartilage Tissue Fabrication Using Three-dimensional Inkjet Printing Technology

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and regenerative medicine. With digital control&#160;cells, scaffolds, and growth factors can be precisely deposited to the desired two-dimensional (2D) and three-dimensional (3D) locations rapidly. Therefore, this technology is an ideal approach to fabricate tissues mimicking their native anatomic structures. In order to engineer cartilage with native zonal organization, extracellular matrix composition (ECM), and mechanical properties, we developed a bioprinting platform using a commercial inkjet printer with simultaneous photopolymerization capable for 3D cartilage tissue engineering. Human chondrocytes suspended in poly(ethylene glycol) diacrylate (PEGDA) were printed for 3D neocartilage construction via layer-by-layer assembly. The printed cells were fixed at their original deposited positions, supported by the surrounding scaffold in simultaneous photopolymerization. The mechanical properties of the printed tissue were similar to the native cartilage. Compared to conventional tissue fabrication, which requires longer UV exposure, the viability of the printed cells with simultaneous photopolymerization was significantly higher. Printed neocartilage demonstrated excellent glycosaminoglycan (GAG) and collagen type II production, which was consistent with gene expression. Therefore, this platform is ideal for accurate cell distribution and arrangement for anatomic tissue engineering.

The methods described in this paper show how to convert a commercial inkjet printer into a bioprinter with simultaneous UV polymerization. The printer is capable of constructing 3D tissue structure with cells and biomaterials. The study demonstrated here constructed a 3D neocartilage.

Bioprinting, which is based on thermal inkjet printing, is one of the most attractive enabling technologies in the field of tissue engineering and ...

Analysis of Cell Migration within a Three-dimensional Collagen Matrix

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic development, wound healing, and immune responses. However, cell migration is also a key mechanism in cancer enabling these cancer cells to detach from the primary tumor to start metastatic spreading. Within the past years various cell migration assays have been developed to analyze the migratory behavior of different cell types. Because the locomotory behavior of cells markedly differs between a two-dimensional (2D) and three-dimensional (3D) environment it can be assumed that the analysis of the migration of cells that are embedded within a 3D environment would yield in more significant cell migration data. The advantage of the described 3D collagen matrix migration assay is that cells are embedded within a physiological 3D network of collagen fibers representing the major component of the extracellular matrix. Due to time-lapse video microscopy real cell migration is measured allowing the determination of several migration parameters as well as their alterations in response to pro-migratory factors or inhibitors. Various cell types could be analyzed using this technique, including lymphocytes/leukocytes, stem cells, and tumor cells. Likewise, also cell clusters or spheroids could be embedded within the collagen matrix concomitant with analysis of the emigration of single cells from the cell cluster/ spheroid into the collagen lattice. We conclude that the 3D collagen matrix migration assay is a versatile method to analyze the migration of cells within a physiological-like 3D environment.

Cell migration is a biological phenomenon that is involved in a plethora of physiological, such as wound healing and immune responses, and pathophysiological processes, like cancer. The 3D-collagen matrix migration assay is a versatile tool to analyze the migratory properties of different cell types within in a 3D physiological-like environment.

The ability to migrate is a hallmark of various cell types and plays a crucial role in several physiological processes, including&#160;embryonic ...

Engineering Three-dimensional Epithelial Tissues Embedded within Extracellular Matrix

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, which is regulated by a variety of soluble and physical signals in the microenvironment. Described here is a method created to study the process of branching morphogenesis by forming engineered three-dimensional (3D) epithelial tissues of defined shape and size that are completely embedded within an extracellular matrix (ECM). This method enables the formation of arrays of identical tissues and enables the control of a variety of environmental factors, including tissue geometry, spacing, and ECM composition. This method can also be combined with widely used techniques such as traction force microscopy (TFM) to gain more information about the interactions between cells and their surrounding ECM. The protocol can be used to investigate a variety of cell and tissue processes beyond branching morphogenesis, including cancer invasion.

This manuscript describes a soft lithography-based technique to engineer uniform arrays of three-dimensional (3D) epithelial tissues of defined geometry surrounded by extracellular matrix. This method is amenable to a wide variety of cell types and experimental contexts and allows for high-throughput screening of identical replicates.

The architecture of branched organs such as the lungs, kidneys, and mammary glands arises through the developmental process of branching morphogenesis, ...

Fabrication of Three-dimensional Paper-based Microfluidic Devices for Immunoassays

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and directed within a layer of paper. Moreover, stacking multiple layers of patterned paper creates sophisticated three-dimensional microfluidic networks that can support the development of analytical and bioanalytical assays. Paper-based microfluidic devices are inexpensive, portable, easy to use, and require no external equipment to operate. As a result, they hold great promise as a platform for point-of-care diagnostics. In order to properly evaluate the utility and analytical performance of paper-based devices, suitable methods must be developed to ensure their manufacture is reproducible and at a scale that is appropriate for laboratory settings. In this manuscript, a method to fabricate a general device architecture that can be used for paper-based immunoassays is described. We use a form of additive manufacturing (multi-layer lamination) to prepare devices that comprise multiple layers of patterned paper and patterned adhesive. In addition to demonstrating the proper use of these three-dimensional paper-based microfluidic devices with an immunoassay for human chorionic gonadotropin (hCG), errors in the manufacturing process that may result in device failures are discussed. We expect this approach to manufacturing paper-based devices will find broad utility in the development of analytical applications designed specifically for limited-resource settings.

We detail a method to fabricate three-dimensional paper-based microfluidic devices for use in the development of immunoassays. Our approach to device assembly is a type of multilayer, additive manufacturing. We demonstrate a sandwich immunoassay to provide representative results for these types of paper-based devices.

Paper wicks fluids autonomously due to capillary action. By patterning paper with hydrophobic barriers, the transport of fluids can be controlled and ...

Switchable Acoustic and Optical Resolution Photoacoustic Microscopy for In Vivo Small-animal Blood Vasculature Imaging

Photoacoustic microscopy (PAM) is a fast-growing invivo imaging modality that combines both optics and ultrasound, providing penetration beyond the optical mean free path (~1 mm in skin) with high resolution. By combining optical absorption contrast with the high spatial resolution of ultrasound in a single modality, this technique can penetrate deep tissues. Photoacoustic microscopy systems can have either a low acoustic resolution and probe deeply or a high optical resolution and probe shallowly. It is challenging to achieve high spatial resolution and large depth penetration with a single system. This work presents an AR-OR-PAM system capable of both high-resolution imaging at shallow depths and low-resolution deep-tissue imaging of the same sample in vivo. A lateral resolution of 4 &#181;m with 1.4 mm imaging depth using optical focusing and a lateral resolution of 45 &#181;m with 7.8 mm imaging depth using acoustic focusing were successfully demonstrated using the combined system. Here,&#160;in vivo&#160;small-animal blood vasculature imaging is performed to demonstrate its biological imaging capability.

Switchable Acoustic and Optical Resolution Photoacoustic Microscopy for In Vivo Small-animal Blood Vasculature Imaging

Here a switchable acoustic resolution (AR) and optical resolution (OR) photoacoustic microscopy (AR-OR-PAM) system capable of both high resolution imaging at shallow depth and low resolution deep tissue imaging on the same sample in vivo is demonstrated.

Photoacoustic microscopy (PAM) is a fast-growing invivo imaging modality that combines both optics and ultrasound, providing penetration beyond the ...

Characterizing Cell Migration Within Three-dimensional In Vitro Wound Environments

Currently, most in vitro models of wound healing, such as well-established scratch assays, involve studying cell migration and wound closure on two-dimensional surfaces. However, the physiological environment in which in vivo wound healing takes place is three-dimensional rather than two-dimensional. It is becoming increasingly clear that cell behavior differs greatly in two-dimensional vs. three-dimensional environments; therefore, there is a need for more physiologically relevant in vitro models for studying cell migration behaviors in wound closure. The method described herein allows for the study of cell migration in a three-dimensional model that better reflects physiological conditions than previously established two-dimensional scratch assays. The purpose of this model is to evaluate cell outgrowth via the examination of cell migration away from a spheroid body embedded within a fibrin matrix in the presence of pro- or anti-migratory factors. Using this method, cell outgrowth from the spheroid body in a three-dimensional matrix can be observed and is easily quantifiable over time via brightfield microscopy and analysis of spheroid body area. The effect of pro-migratory and/or inhibitory factors on cell migration can also be evaluated in this system. This method provides researchers with a simple method of analyzing cell migration in three-dimensional wound associated matrices in vitro, thus increasing the relevance of in vitro cell studies prior to the use of in vivo animal models.

Characterizing Cell Migration Within Three-dimensional In Vitro Wound Environments

The goal of this protocol is to evaluate the effect of pro- and anti-migratory factors on cell migration within a three-dimensional fibrin matrix.

Currently, most in vitro models of wound healing, such as well-established scratch assays, involve studying cell migration and wound closure on ...

Culturing Mammalian Cells in Three-dimensional Peptide Scaffolds

A useful technique for culturing cells in a self-assembling nanofiber three-dimensional (3D) scaffold is described. This culture system recreates an environment that closely mimics the structural features of non-polarized tissue. Furthermore, the particular intrinsic nanofiber structure of the scaffold makes it transparent to visual light, which allows for easy visualization of the sample under microscopy. This advantage was largely used to study cell migration, organization, proliferation, and differentiation and thus any development of their particular cellular function by staining with specific dyes or probes. Furthermore, in this work, we describe the good performance of this system to easily study the redifferentiation of expanded human articular chondrocytes into cartilaginous tissue. Cells were encapsulated into self-assembling peptide scaffolds and cultured under specific conditions to promote chondrogenesis. Three-dimensional cultures showed good viability during the 4 weeks of the experiment. As expected, samples cultured with chondrogenic inducers (compared to non-induced controls) stained strongly positive for toluidine blue (which stains glycosaminoglycans (GAGs) that are highly present in cartilage extracellular matrix) and expressed specific molecular markers, including collagen type I, II and X, according to Western Blot analysis. This protocol is easy to perform and can be used at research laboratories, industries and for educational purposes in laboratory courses.

Here, we present a protocol to obtain 3D-culture systems in self-assembling peptide scaffolds to promote the differentiation of dedifferentiated human articular chondrocytes into cartilage-like tissue.

A useful technique for culturing cells in a self-assembling nanofiber three-dimensional (3D) scaffold is described. This culture system recreates an ...