We are grateful to Dr. Kouki Hikosaka (Tohoku University) for stimulating discussions, assistance with a work space, and instruments for experiments. The work was supported in part by KAKENHI [grant numbers 18K05592, 18J40098] and Naito Foundation.

1. Cultivation of Arabidopsis plants
<ol>
	<li>Soak Arabidopsis thaliana seeds in sterilized deionized water in a microtube, and incubate for 2 days at 4 &#176;C in the dark.</li>
	<li>Place approximately four of the imbibed, cold-treated seeds onto the soil surface using a micropipette. Incubate the planted pots in a growth chamber with a 16 h light (120 &#956;mol photons m&#8211;2 s&#8211;1) and 8 h dark period at 22 &#176;C and 20 &#176;C, respectively.</li>
	<li>Grow one plant per pot by thinning other seedlings after germination. Prepare at least five pots. Incubate the plants in the growth chamber for an additional 4 weeks. Three plants are used for the experiments.</li>
	<li>Use the youngest, fully-opened mature leaf for photosynthesis measurements.</li>
</ol>
2. Setting up the sample stage, photosynthetic instruments, and light source
NOTE: For this protocol, a custom-built sample stage was used for fixing leaves and detection probes (Figure 1).
<ol>
	<li>Attach a 10 cm2 steel plate with a 1 cm diameter hole onto the custom-made sample stage. The hole size in this plate can be changed to accommodate different leaf samples or plant species. The stage has a clip for fixing the detection probes and an adjuster to adjust the distance between the probes and the leaf sample.</li>
	<li>Prepare thin fiber probes for measuring chlorophyll fluorescence and leaf reflectance. These thin fiber probes will be positioned closely so that they measure signals from the same leaf position. 
	NOTE: A PAM chlorophyll fluorometer and a spectral radiometer were adapted for signal detection of chlorophyll a fluorescence and leaf reflectance, respectively. Both instruments use thin fiber probes with diameters of 1 mm and 2 mm, respectively.</li>
	<li>Fit these two probes tightly together and wrap them with plastic tape.</li>
	<li>Clip the taped probes onto the sample stage using a coaxial lens holder (see Figure 1), and position them vertical to the leaf surface. 
	NOTE: The field of view of the fiber optic cable in the spectral radiometer is &#945; = 25&#176;. In this method, the distance between the fiber probe tips and leaf surface is shorter than 1 cm. Therefore, the measuring leaf area is almost the same as that of the fiber.</li>
	<li>Attach a biforked light guide made of glass fibers to the halogen light source and irradiate the sample stage from both directions at angles of approximately 45&#176;. 
	NOTE: A halogen lamp, which is close to the wavelength distribution of natural sunlight, is used as actinic light for inducing photosynthesis. The halogen light source was adapted with a built-in cold filter, which removes long wavelengths from near infrared, to prevent increases in leaf surface temperature (&#955; = 400 to 800 nm).</li>
	<li>Adjust the light source so that light uniformly illuminates the sample stage without casting shadows.</li>
</ol>
3. Setting up simultaneous measurements of leaf reflectance and chlorophyll fluorescence
NOTE: All steps are performed in the dark room to avoid the detection of light other than actinic light. A weak-green light (e.g., green-cellophaned light) should be turned off before the actual measurements.
<ol>
	<li>Measuring the distance between the leaf sample and the probes on the sample stage.
	<ol>
		<li>Place a test leaf on the leaf holder of the sample stage in the dark. Press the leaf against a steel plate on the stage (black square in Figure 1).</li>
		<li>Turn on the PAM and irradiate the leaf sample with a measuring light. The values of the chlorophyll fluorescence intensities are confirmed using PAM controlling software (see Table of Materials).</li>
		<li>Move the adjuster so that the fluorescence intensity measures approximately 100. Measure the distance between the probe and the leaf. Fix the adjuster, and record the value of the distance on the adjuster.</li>
		<li>Turn off the measuring light. Remove the test leaf.</li>
	</ol>
	</li>
	<li>Measuring the irradiance intensities of the actinic light 
	NOTE: To observe light-dependent photosynthetic behaviors, actinic light of varying intensity is used to irradiate the leaf sample.
	<ol>
		<li>Set a light quantum meter at the position where the sample leaf would be placed.</li>
		<li>Irradiate light from the halogen light source and measure the intensity.</li>
		<li>Determine which positions of the light source dial would generate intensities of 30, 60, 120, 240, and 480 &#956;mol photons m&#8211;2 s&#8211;1. 
		NOTE: Arabidopsis plants are grown under 120 &#956;mol photons m&#8211;2 s&#8211;1; therefore, the irradiance intensities of the actinic light are selected to provide a range of small and large intensities.</li>
		<li>Mark each irradiance intensity on the dial.</li>
	</ol>
	</li>
	<li>Measure a reflection standard. 
	NOTE: A reflection standard is required to calculate the leaf reflectance ratio at each irradiance intensity.
	<ol>
		<li>Place a white plate as a reflectance standard at the position of the leaf sample.</li>
		<li>Turn on a spectral radiometer. The reflectance signal is shown by the spectral radiometer controlling software. At this time, there are no spectral data because there is no irradiating light.</li>
		<li>Turn on the halogen lamp to irradiate with 480 &#956;mol photons m&#8211;2 s&#8211;1, the highest irradiance intensity in this test.</li>
		<li>Adjust the detection strength of the radiometer to avoid saturation.</li>
		<li>Record spectral reflectance between 450&#8211;850 nm at 1 nm intervals under illumination with 30, 60, 120, 240, and 480 &#956;mol photons m&#8211;2 s&#8211;1. 
		NOTE: A baseline electrical signal (dark current) is corrected and subtracted at every spectral measurement.</li>
	</ol>
	</li>
</ol>
4. Simultaneous measurements of leaf reflectance and chlorophyll a fluorescence, and calculation of photosynthetic parameters
<ol>
	<li>Set a plant at the leaf sample position.
	<ol>
		<li>Transfer the Arabidopsis plant from the grown chamber to the controlled dark room with the same temperature and humidity as that of the growth chamber.</li>
		<li>Incubate the plant for 1 h in the dark at 22 &#176;C to dissipate electrons from the PSII reaction center and to relax of non-photochemical quenching.</li>
		<li>Place the dark-adapted whole plant on a lab jack under the sample stage (Figure 1).</li>
		<li>Fix the sample leaf to the leaf holder so that the leaf surface is perpendicular to the detection probes.</li>
	</ol>
	</li>
	<li>Measurement of the maximum quantum yield of PSII.
	<ol>
		<li>Turn on the PAM and start recording the curve. This value is called 0.</li>
		<li>Turn on the measuring light, and wait approximately 30 s for the curve to respond. This value is called F0.</li>
		<li>Give a saturated pulse of 4000 &#956;mol photons m&#8211;2 s&#8211;1 for 0.8 s from the PAM.</li>
		<li>Obtain the highest value of the spike in the curve with increased fluorescence intensity. This value is called FM.</li>
		<li>Calculate the maximum quantum yield of PSII in the dark (FV/FM), using the following equation. 
		FV/FM = (FM &#8211; F0) / FM</li>
	</ol>
	</li>
	<li>Measurement of photosynthetic behaviors at steady state.
	<ol>
		<li>Turn on the halogen lamp as the external light source with the measuring light on after recording FM (see 4.2.4). First, irradiate the leaf sample with the weakest light (30 &#956;mol photons m&#8211;2 s&#8211;1).</li>
		<li>Turn on the spectral radiometer at the same time to monitor the leaf reflectance.</li>
		<li>Wait for 20 min or longer for the photosynthetic reaction to reach steady state under the light conditions. The fluorescence intensity of the steady state is called Fs.</li>
		<li>Supply a saturating pulse at intervals of 1 min during illumination with the actinic light. The maximum fluorescence value achieved under the pulsed light is called FM&#8242;.</li>
		<li>Record the data of FM&#8242; at 20 min after turning on the actinic light.</li>
		<li>Take leaf reflectance data by averaging 10 scans at an optimized integration time, with a dark current subtraction.</li>
	</ol>
	</li>
	<li>Calculation of photosynthetic parameters at steady state.
	<ol>
		<li>Calculate the quantum yields of PSII photochemistry (&#934;PSII), which can be estimated by irradiating with saturated pulses under actinic light, using the following equation. 
		&#934;PSII = (FM&#8242;- FS) / FM&#8242;</li>
		<li>Estimate the linear electron flux (LEF) from the PSII reaction center as follows 4. 
		LEF = The irradiance intensity of the actinic light &#215; &#934;PSII &#215; 0.5 &#215; 0.84</li>
		<li>Calculate the NPQ, which can be expressed the thermal dissipation, using the following equation. 
		NPQ = (FM - FM&#8242;) / FM&#8242; 
		NOTE: Light energy is primarily consumed by photosynthesis reactions. However, when plants absorb more light energy than energy consumed by photosynthesis, the mechanisms for the thermal dissipation are induced to avoid the excess energy.</li>
		<li>Using the spectral data acquired with the radiometer under the same light conditions, calculate the leaf reflectance ratio as follows. 
		Reflectance ratio = Rleaf / Rstandard</li>
		<li>Calculate PRI from 531 nm and 570 nm as follows. These two wavelengths are extracted from the reflectance ratio. 
		PRI = (R531&#8211;R570) / (R531+R570) 
		NOTE: R is a reflectance.</li>
	</ol>
	</li>
	<li>Measuring of the relaxation kinetics of non-photochemical quenching.
	<ol>
		<li>Turn off the actinic light after acquiring Fs and leaf reflectance.</li>
		<li>Monitor chlorophyll fluorescence by PAM for 10 min after turning off light.</li>
		<li>Provide a saturating pulse at intervals of 1 min during the dark relaxation. The maximum fluorescence value induced by the saturation pulse under the dark is called FM&#8242;&#8242;. Obtain ten of FM&#8242;&#8242; in one test.</li>
		<li>Save the data of FM&#8242;&#8242; at 2 min and 10 min after turning off the actinic light.</li>
		<li>Turn the actinic light on set to the next irradiance intensity, 60 &#956; mol photons m&#8211;2 s&#8211;1.</li>
		<li>Repeat a light adaptation for 20 min and a dark relaxation for 10 min with pulsing saturation light at intervals of 1 min. Record all data as described above. Repeat all steps and measurements using irradiation at 120, 240, and 480 &#956; mol photons m&#8211;2 s&#8211;1.</li>
	</ol>
	</li>
	<li>Calculation of parameters of the non-photochemical quenching from the relaxation kinetics. 
	NOTE: Light-dependent induction of NPQ is relaxed by turning off the light source13. It is possible to fractionate each NPQ function by adjusting the relaxation timescales.
	<ol>
		<li>Estimate the qE (energy-dependent quenching) fraction using FM&#8242;&#8242; after 2-min dark adaptation. 
		qE = (FM2m&#8242;&#8242;&#8211;FM&#8242;) / FM&#8242; 
		NOTE: The qE fraction is rapidly reversed within 1&#8211;2 min. This fraction includes mainly PsbS protonation and part of xanthophyll conversion, which depend on light-induced &#916;pH across the thylakoid membrane13. Both are reversible on breakdown of the gradient.</li>
		<li>Calculate the qZ (zeaxanthin dependent quenching) fraction using FM&#8242;&#8242; after 10-min dark adaptation. 
		qZ = (FM10m&#8242;&#8242;&#8211;FM&#8242;) / FM&#8242; 
		NOTE: A relaxation kinetics of NPQ at approximately 10 min after actinic light off reflects a xanthophyll cycle14. Most of the xanthophyll conversion is reversed at longer timescales of approximately 10 min (qZ) because the conversion requires a VDE (violaxanthin de-epoxidase) enzymatic reaction. The fraction is also relaxed by breakdown of the &#916;pH across the thylakoid membrane.</li>
		<li>Calculate the qI (photoinhibitory state) as follows. 
		qI = (FM&#8211;FM10m&#8242;&#8242;) / FM&#8242; 
		NOTE: The slowest recovery among NPQ fractions is thought to be photodamage of PSII (indicating D1 turnover). This fraction of the photoinhibitory state (qI), which does not recover by 10 min15.</li>
	</ol>
	</li>
</ol>

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The author has nothing to disclose.

In this study, we obtained additional evidence to show that PRI represents xanthophyll pigments by simultaneously measuring chlorophyll fluorescence and leaf reflectance.
A halogen light, which has wavelengths similar to sunlight, was adapted for use as an actinic light source to activate photosynthesis. We initially used a white LED light source to avoid thermal damage of the leaf surface, but this produced slow dark relaxation kinetics and exceptionally high qI (photoinhibitory quenching), p...

Leaf reflectance is used to remotely sense vegetation indices that reflect photosynthesis or traits in plants1,2. The normalized difference vegetation index (NDVI), which is based on infrared reflection signals, is one of the most widely known vegetation indices for the detection of chlorophyll-related properties, and it is used in the ecology and agricultural sciences as an indicator of environmental responses in trees or crops3. In field studies, although many parameters (e.g., chlorophyll index (CI), water index (WI), etc.) have been developed and used, few detailed verifications of what these parameters directly (or indirectly) detect have been performed using mutants.
Pulse-amplitude modulation (PAM) analysis of chlorophyll fluorescence is an effective method to measure photosynthetic reactions and processes involved in photosystem II (PSII)4. Chlorophyll fluorescence can be detected with a camera and used for screening photosynthesis mutants5. However, camera detection of chlorophyll fluorescence requires complex protocols such as dark treatment or light saturation pulses, which are difficult to implement in field studies.
Leaf absorbed solar light energy is mainly consumed by photosynthetic reactions. By contrast, the absorption of excess light energy can generate reactive oxygen species, which causes damage to photosynthetic molecules. The excess light energy must be dissipated as heat through non-photochemical quenching (NPQ) mechanisms6. The photochemical reflectance index (PRI), which reflects light-dependent changes in leaf reflectance parameters, is derived from narrow-band reflectance at 531 and 570 nm (reference wavelength)7,8. It is reported to correlate with NPQ in chlorophyll fluorescence analysis9. However, since NPQ is a composite parameter that includes the xanthophyll cycle, state tradition, and photoinhibition, detailed validation is required to understand what the PRI parameter measures. We have focused on the xanthophyll cycle, a thermal dissipation system involving the de-epoxidation of xanthophyll pigments (violaxanthin to antheraxanthin and zeaxanthin) and a main component of NPQ because correlations between PRI and conversion of these pigments has been reported in previous studies8.
Many photosynthesis-related mutants have been isolated and identified in Arabidopsis. The npq1 mutant does not accumulate zeaxanthin because it carries a mutation in violaxanthin de-epoxidase (VDE), which catalyzes the conversion of violaxanthin to zeaxanthin10. To establish whether PRI only detects changes in xanthophyll pigments, we simultaneously measured PRI and chlorophyll fluorescence in the same leaf area in npq1 and the wild-type and then dissected NPQ at varying time scales of dark relaxation to extract the xanthophyll-related component11. These simultaneous measurements provide a valuable technique for the assignment of vegetation indices. Furthermore, since PRI correlates with gross primary productivity (GPP), the ability to assign PRI precisely to one component has important applications in ecology12.

<table>
	<tbody>
		<tr>
			<td>Halogen light source</td>
			<td>OptoSigma</td>
			<td>SHLA-150</td>
			<td></td>
		</tr>
		<tr>
			<td>Light quantum meter</td>
			<td>LI-COR</td>
			<td>LI-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>PAM chlorophyll fluorometer</td>
			<td>Walz</td>
			<td>JUNIOR-PAM</td>
			<td></td>
		</tr>
		<tr>
			<td>PAM controliing software</td>
			<td>Walz</td>
			<td>WinControl-3.27</td>
			<td></td>
		</tr>
		<tr>
			<td>Reflectance standard</td>
			<td>Labsphere, Inc.</td>
			<td>SRT-99-050</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectral radiometer</td>
			<td>ADS Inc.</td>
			<td>Field Spec3</td>
			<td></td>
		</tr>
		<tr>
			<td>Spectral radiometer controlling software</td>
			<td>ADS Inc.</td>
			<td>RS3</td>
			<td></td>
		</tr>
	</tbody>
</table>

evaluation of photosynthetic behaviors by simultaneous measurements of leaf reflectance and chlorophyll fluorescence analyses

Chlorophyll a fluorescence analysis is widely used to measure photosynthetic behaviors in intact plants, and has resulted in the development of many parameters that efficiently measure photosynthesis. Leaf reflectance analysis provides several vegetation indices in ecology and agriculture, including the photochemical reflectance index (PRI), which can be used as an indicator of thermal energy dissipation during photosynthesis because it correlates with non-photochemical quenching (NPQ). However, since NPQ is a composite parameter, its validation is required to understand the nature of the PRI parameter. To obtain physiological evidence for evaluation of the PRI parameter, we simultaneously measured chlorophyll fluorescence and leaf reflectance in xanthophyll cycle defective mutant (npq1) and wild-type Arabidopsis plants. Additionally, the qZ parameter, which likely reflects the xanthophyll cycle, was extracted from the results of chlorophyll fluorescence analysis by monitoring relaxation kinetics of NPQ after switching the light off. These simultaneous measurements were carried out using a pulse-amplitude modulation (PAM) chlorophyll fluorometer and a spectral radiometer. The fiber probes from both instruments were positioned close to each other to detect signals from the same leaf position. An external light source was used to activate photosynthesis, and the measuring lights and saturated light were provided from the PAM instrument. This experimental system enabled us to monitor light-dependent PRI in the intact plant and revealed that light-dependent changes in PRI differ significantly between the wild type and npq1 mutant. Furthermore, PRI was strongly correlated with qZ, meaning that qZ reflects the xanthophyll cycle. Together, these measurements demonstrated that simultaneous measurement of leaf reflectance and chlorophyll fluorescence is a valid approach for parameter evaluation.

Figure 1 presents a schematic diagram of the experimental set up for simultaneously measuring chlorophyll fluorescence and leaf reflectance. The fiber probes of the PAM and spectral radiometer were set perpendicularly to the leaf surface at the leaf holder on the custom-made sample stage, and a halogen lamp was used for actinic light irradiation from both left and right directions without casting any shadows. The PAM and leaf reflectance signals were detected...

Watch this Scientific Journal Video about Evaluation of Photosynthetic Behaviors by Simultaneous Measurements of Leaf Reflectance and Chlorophyll Fluorescence Analyses at JoVE.com

Evaluation of Photosynthetic Behaviors by Simultaneous Measurements of Leaf Reflectance and Chlorophyll Fluorescence Analyses

We describe a new technical approach to study photosynthetic responses in higher plants involving simultaneous measurements of chlorophyll a fluorescence and leaf reflectance using a PAM and a spectral radiometer for the detection of signals from the same leaf area in Arabidopsis.

Chlorophyll a fluorescence analysis is widely used to measure photosynthetic behaviors in intact plants, and has resulted in the development of many ...

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12.2K Views.  Tohoku University. We describe a new technical approach to study photosynthetic responses in higher plants involving simultaneous measurements of chlorophyll a fluorescence and leaf reflectance using a PAM and a spectral radiometer for the detection of signals from the same leaf area in Arabidopsis.

Video: Evaluation of Photosynthetic Behaviors by Simultaneous Measurements of Leaf Reflectance and Chlorophyll Fluorescence Analyses

Thermodynamics of Membrane Protein Folding Measured by Fluorescence Spectroscopy

Membrane protein folding is an emerging topic with both fundamental and health-related significance. The abundance of membrane proteins in cells underlies the need for comprehensive study of the folding of this ubiquitous family of proteins. Additionally, advances in our ability to characterize diseases associated with misfolded proteins have motivated significant experimental and theoretical efforts in the field of protein folding. Rapid progress in this important field is unfortunately hindered by the inherent challenges associated with membrane proteins and the complexity of the folding mechanism. Here, we outline an experimental procedure for measuring the thermodynamic property of the Gibbs free energy of unfolding in the absence of denaturant, &Delta;G&deg;H2O, for a representative integral membrane protein from E. coli. This protocol focuses on the application of fluorescence spectroscopy to determine equilibrium populations of folded and unfolded states as a function of denaturant concentration. Experimental considerations for the preparation of synthetic lipid vesicles as well as key steps in the data analysis procedure are highlighted. This technique is versatile and may be pursued with different types of denaturant, including temperature and pH, as well as in various folding environments of lipids and micelles. The current protocol is one that can be generalized to any membrane or soluble protein that meets the set of criteria discussed below.

This video article details the experimental procedure for obtaining the Gibbs free energy of membrane protein folding by tryptophan fluorescence.

Membrane protein folding is an emerging topic with both fundamental and health-related significance.  The abundance of membrane proteins in cells ...

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, ...

Evaluation of Biomaterials for Bladder Augmentation using Cystometric Analyses in Various Rodent Models

Renal function and continence of urine are critically dependent on the proper function of the urinary bladder, which stores urine at low pressure and expels it with a precisely orchestrated contraction. A number of congenital and acquired urological anomalies including posterior urethral valves, benign prostatic hyperplasia, and neurogenic bladder secondary to spina bifida/spinal cord injury can result in pathologic tissue remodeling leading to impaired compliance and reduced capacity1. Functional or anatomical obstruction of the urinary tract is frequently associated with these conditions, and can lead to urinary incontinence and kidney damage from increased storage and voiding pressures2. Surgical implantation of gastrointestinal segments to expand organ capacity and reduce intravesical pressures represents the primary surgical treatment option for these disorders when medical management fails3. However, this approach is hampered by the limitation of available donor tissue, and is associated with significant complications including chronic urinary tract infection, metabolic perturbation, urinary stone formation, and secondary malignancy4,5.
Current research in bladder tissue engineering is heavily focused on identifying biomaterial configurations which can support regeneration of tissues at defect sites. Conventional 3-D scaffolds derived from natural and synthetic polymers such as small intestinal submucosa and poly-glycolic acid have shown some short-term success in supporting urothelial and smooth muscle regeneration as well as facilitating increased organ storage capacity in both animal models and in the clinic6,7. However, deficiencies in scaffold mechanical integrity and biocompatibility often result in deleterious fibrosis8, graft contracture9, and calcification10, thus increasing the risk of implant failure and need for secondary surgical procedures. In addition, restoration of normal voiding characteristics utilizing standard biomaterial constructs for augmentation cystoplasty has yet to be achieved, and therefore research and development of novel matrices which can fulfill this role is needed.
In order to successfully develop and evaluate optimal biomaterials for clinical bladder augmentation, efficacy research must first be performed in standardized animal models using detailed surgical methods and functional outcome assessments. We have previously reported the use of a bladder augmentation model in mice to determine the potential of silk fibroin-based scaffolds to mediate tissue regeneration and functional voiding characteristics.11,12 Cystometric analyses of this model have shown that variations in structural and mechanical implant properties can influence the resulting urodynamic features of the tissue engineered bladders11,12. Positive correlations between the degree of matrix-mediated tissue regeneration determined histologically and functional compliance and capacity evaluated by cystometry were demonstrated in this model11,12. These results therefore suggest that functional evaluations of biomaterial configurations in rodent bladder augmentation systems may be a useful format for assessing scaffold properties and establishing in vivo feasibility prior to large animal studies and clinical deployment. In the current study, we will present various surgical stages of bladder augmentation in both mice and rats using silk scaffolds and demonstrate techniques for awake and anesthetized cystometry.

Surgical stages of bladder augmentation are described using 3-D scaffolds in murine and rat models. To test the efficacy of biomaterial configurations for use in bladder augmentation, techniques for both awake and anesthetized cystometry are presented.

Renal function and continence of urine are critically dependent on the proper function of the urinary bladder, which stores urine at low pressure and ...

Evaluation of Polymeric Gene Delivery Nanoparticles by Nanoparticle Tracking Analysis and High-throughput Flow Cytometry

Non-viral gene delivery using polymeric nanoparticles has emerged as an attractive approach for gene therapy to treat genetic diseases1 and as a technology for regenerative medicine2. Unlike viruses, which have significant safety issues, polymeric nanoparticles can be designed to be non-toxic, non-immunogenic, non-mutagenic, easier to synthesize, chemically versatile, capable of carrying larger nucleic acid cargo and biodegradable and/or environmentally responsive. Cationic polymers self-assemble with negatively charged DNA via electrostatic interaction to form complexes on the order of 100 nm that are commonly termed polymeric nanoparticles. Examples of biomaterials used to form nanoscale polycationic gene delivery nanoparticles include polylysine, polyphosphoesters, poly(amidoamines)s and polyethylenimine (PEI), which is a non-degradable off-the-shelf cationic polymer commonly used for nucleic acid delivery1,3 . Poly(beta-amino ester)s (PBAEs) are a newer class of cationic polymers4 that are hydrolytically degradable5,6 and have been shown to be effective at gene delivery to hard-to-transfect cell types such as human retinal endothelial cells (HRECs)7, mouse mammary epithelial cells8, human brain cancer cells9 and macrovascular (human umbilical vein, HUVECs) endothelial cells10.
A new protocol to characterize polymeric nanoparticles utilizing nanoparticle tracking analysis (NTA) is described. In this approach, both the particle size distribution and the distribution of the number of plasmids per particle are obtained11. In addition, a high-throughput 96-well plate transfection assay for rapid screening of the transfection efficacy of polymeric nanoparticles is presented. In this protocol, poly(beta-amino ester)s (PBAEs) are used as model polymers and human retinal endothelial cells (HRECs) are used as model human cells. This protocol can be easily adapted to evaluate any polymeric nanoparticle and any cell type of interest in a multi-well plate format.

A protocol for nanoparticle tracking analysis (NTA) and high-throughput flow cytometry to evaluate polymeric gene delivery nanoparticles is described. NTA is utilized to characterize the nanoparticle particle size distribution and the plasmid per particle distribution. High-throughput flow cytometry enables quantitative transfection efficacy evaluation for a library of gene delivery biomaterials.

Non-viral gene delivery using polymeric nanoparticles has emerged as an attractive approach for gene therapy to treat genetic diseases1 and as a ...

Aplysia Ganglia Preparation for Electrophysiological and Molecular Analyses of Single Neurons

A major challenge in neurobiology is to understand the molecular underpinnings of neural circuitry that govern a specific behavior. Once the specific molecular mechanisms are identified, new therapeutic strategies can be developed to treat abnormalities in specific behaviors caused by degenerative diseases or aging of the nervous system. The marine snail Aplysia californica is well suited for the investigations of cellular and molecular basis of behavior because neural circuitry underlying a specific behavior could be easily determined and the individual components of the circuitry could be easily manipulated. These advantages of Aplysia have led to several fundamental discoveries of neurobiology of learning and memory. Here we describe a preparation of the Aplysia nervous system for the electrophysiological and molecular analyses of individual neurons. Briefly, ganglion dissected from the nervous system is exposed to protease to remove the ganglion sheath such that neurons are exposed but retain neuronal activity as in the intact animal. This preparation is used to carry out electrophysiological measurements of single or multiple neurons. Importantly, following the recording using a simple methodology, the neurons could be isolated directly from the ganglia for gene expression analysis. These protocols were used to carry out simultaneous electrophysiological recordings from L7 and R15 neurons, study their response to acetylcholine and quantitating expression of CREB1 gene in isolated single L7, L11, R15, and R2 neurons of Aplysia.

Aplysia Ganglia Preparation for Electrophysiological and Molecular Analyses of Single Neurons

Marine snail Aplysia californica has been widely used as a neurobiology model for the studies on cellular and molecular basis of behavior. Here a methodology is described for exploring the nervous system of Aplysia for the electrophysiological and molecular analyses of single neurons of identified neural circuitry.

A major challenge in neurobiology is to understand the molecular underpinnings of neural circuitry that govern a specific behavior. Once the specific ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

3D Orbital Tracking in a Modified Two-photon Microscope: An Application to the Tracking of Intracellular Vesicles

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a modified two-photon microscope1. As opposed to conventional approaches (raster scan or wide field based on a stack of frames), the 3D orbital tracking allows to localize and follow with a high spatial (10 nm accuracy) and temporal resolution (50 Hz frequency response) the 3D displacement of a moving fluorescent particle on length-scales of hundreds of microns2. The method is based on a feedback algorithm that controls the hardware of a two-photon laser scanning microscope in order to perform a circular orbit around the object to be tracked: the feedback mechanism will maintain the fluorescent object in the center by controlling the displacement of the scanning beam3-5. To demonstrate the advantages of this technique, we followed a fast moving organelle, the lysosome, within a living cell6,7. Cells were plated according to standard protocols, and stained using a commercially lysosome dye. We discuss briefly the hardware configuration and in more detail the control software, to perform a 3D orbital tracking experiment inside living cells. We discuss in detail the parameters required in order to control the scanning microscope and enable the motion of the beam in a closed orbit around the particle. We conclude by demonstrating how this method can be effectively used to track the fast motion of a labeled lysosome along microtubules in 3D within a live cell. Lysosomes can move with speeds in the range of 0.4-0.5 &#181;m/sec, typically displaying a directed motion along the microtubule network8.

In this video protocol we track - at high speed and in three dimensions - fluorescently labeled lysosomes within living cells, using the orbital tracking method in a modified two-photon microscope.

The objective of this video protocol is to discuss how to perform and analyze a three-dimensional fluorescent orbital particle tracking experiment using a ...

Dissection, MicroCT Scanning and Morphometric Analyses of the Baculum

Modern morphometrics provides powerful methods to quantify size and shape variation. A basic requirement is a list of coordinates that define landmarks; however such coordinates must represent homologous structures across specimens. While many biological objects consist of easily identified landmarks to satisfy the assumption of homology, many lack such structures. One potential solution is to mathematically place semi-landmarks on an object that represent the same morphological region across specimens. Here, we illustrate a recently developed pipeline to mathematically define semi-landmarks from the mouse baculum (penis bone). Our methods should be applicable to a wide range of objects.

Many biological structures lack easily definable landmarks, making it difficult to apply modern morphometric methods. Here we illustrate methods to study the mouse baculum (a bone in the penis), including dissection and microCT scanning, followed by computational methods to define semi-landmarks that are used to quantify size and shape variation.

Modern morphometrics provides powerful methods to quantify size and shape variation. A basic requirement is a list of coordinates that define landmarks; ...

Construction and Setup of a Bench-scale Algal Photosynthetic Bioreactor with Temperature, Light, and pH Monitoring for Kinetic Growth Tests

The optimal design and operation of photosynthetic bioreactors (PBRs) for microalgal cultivation is essential for improving the environmental and economic performance of microalgae-based biofuel production. Models that estimate microalgal growth under different conditions can help to optimize PBR design and operation. To be effective, the growth parameters used in these models must be accurately determined. Algal growth experiments are often constrained by the dynamic nature of the culture environment, and control systems are needed to accurately determine the kinetic parameters. The first step in setting up a controlled batch experiment is live data acquisition and monitoring. This protocol outlines a process for the assembly and operation of a bench-scale photosynthetic bioreactor that can be used to conduct microalgal growth experiments. This protocol describes how to size and assemble a flat-plate, bench-scale PBR from acrylic. It also details how to configure a PBR with continuous pH, light, and temperature monitoring using a data acquisition and control unit, analog sensors, and open-source data acquisition software.

This paper describes the assembly process and operation of a bench-scale photosynthetic bioreactor that can be used, in conjunction with other methods, to estimate pertinent kinetic growth parameters. This system continuously monitors the pH, light, and temperature using sensors, a data acquisition and control unit, and open-source data acquisition software.

The optimal design and operation of photosynthetic bioreactors (PBRs) for microalgal cultivation is essential for improving the environmental and economic ...

Engineering 'Golden' Fluorescence by Selective Pressure Incorporation of Non-canonical Amino Acids and Protein Analysis by Mass Spectrometry and Fluorescence

Fluorescent proteins are fundamental tools for the life sciences, in particular for fluorescence microscopy of living cells. While wild-type and engineered variants of the green fluorescent protein from Aequorea victoria (avGFP) as well as homologs from other species already cover large parts of the optical spectrum, a spectral gap remains in the near-infrared region, for which avGFP-based fluorophores are not available. Red-shifted fluorescent protein (FP) variants would substantially expand the toolkit for spectral unmixing of multiple molecular species, but the naturally occurring red-shifted FPs derived from corals or sea anemones have lower fluorescence quantum yield and inferior photo-stability compared to the avGFP variants. Further manipulation and possible expansion of the chromophore&#39;s conjugated system towards the far-red spectral region is also limited by the repertoire of 20 canonical amino acids prescribed by the genetic code. To overcome these limitations, synthetic biology can achieve further spectral red-shifting via insertion of non-canonical amino acids into the chromophore triad. We describe the application of SPI to engineer avGFP variants with novel spectral properties. Protein expression is performed in a tryptophan-auxotrophic E. coli strain and by supplementing growth media with suitable indole precursors. Inside the cells, these precursors are converted to the corresponding tryptophan analogs and incorporated into proteins by the ribosomal machinery in response to UGG codons. The replacement of Trp-66 in the enhanced &#34;cyan&#34; variant of avGFP (ECFP) by an electron-donating 4-aminotryptophan results in GdFP featuring a 108 nm Stokes shift and a strongly red-shifted emission maximum (574 nm), while being thermodynamically more stable than its predecessor ECFP. Residue-specific incorporation of the non-canonical amino acid is analyzed by mass spectrometry. The spectroscopic properties of GdFP are characterized by time-resolved fluorescence spectroscopy as one of the valuable applications of genetically encoded FPs in life sciences.

Engineering &#039;Golden&#039; Fluorescence by Selective Pressure Incorporation of Non-canonical Amino Acids and Protein Analysis by Mass Spectrometry and Fluorescence

Synthetic biology enables the engineering of proteins with unprecedented properties using the co-translational insertion of non-canonical amino acids. Here, we presented how a spectrally red-shifted variant of a GFP-type fluorophore with novel fluorescence spectroscopic properties, termed &#34;gold&#34; fluorescent protein (GdFP), is produced in E. coli via selective pressure incorporation (SPI).