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Air Sacs

Air Sacs: Specialized respiratory structures found in birds and some other avian species.
Air sacs are part of the respiratory system and play a crucial role in the efficient delivery of oxygen to tissues and the removal of carbon dioxide.
These thin-walled, balloon-like structures connect to the lungs and facilitate the movement of air through the body, allowing for enhanced gas exchange during both inhalation and exhalation.
Air sacs contribute to the unique breathing mechansim of birds, which is adapted for powered flight.
Studying the structure and function of air sacs is important for understanding avian physiology and biology.

Most cited protocols related to «Air Sacs»

1
Zebrafish Phenotypic Screening Protocol
Zebrafish were maintained in accordance with UK Home Office regulations, UK Animals (Scientific Procedures) Act 1986, under project licence 80/2192, which was reviewed by The Wellcome Trust Sanger Institute Ethical Review Committee.
Heterozygous F2 fish were randomly incrossed and upon egg collection F2 adults were fin clipped and kept as isolated breeding pairs. For each family we aimed to phenotype 12 pairs, over 3 weeks of breeding. Each clutch of eggs, which was labelled with the breeding pair ID, was sorted into three 10cm petri dishes of ~50 embryos each. Embryos were incubated at 28.5°C. Previous mutagenesis screens were used as a reference for the phenotyping 27 (link),28 (link). Those phenotypes studied were: day 1 – early patterning defects, early arrest, notochord, eye development, somites, patterning and cell death in the brain; day 2 – cardiac defects, circulation of the blood, pigment (melanocytes), eye and brain development; day 3 – cardiac defects, circulation of the blood, pigment (melanocytes), movement and hatching; day 4 – cardiac defects, movement, pigment (melanocytes) and muscle defects; day 5 – behaviour (hearing, balance, response to touch), swim bladder, pigment (melanocytes, xanthophores and iridophores), distribution of pigment, jaw, skull, axis length, body shape, notochord degeneration, digestive organs (intestinal folds, liver and pancreas), left-right patterning. In the first round of the phenotyping, all phenotypic embryos were discarded. At 5 dpf, >48 phenotypically wild-type embryos were harvested. Embryos were fixed in 100% methanol and stored at −20°C until genotyping was initiated. In the second round, F2s that were heterozygous for a suspected causal mutation were re-crossed. All phenotypes observed in those clutches of embryos were counted, documented and photographed. Phenotypic embryos were fixed in 100% methanol and at 5 dpf 48 phenotypically wild-type embryos were also collected. The first round genotyping results were assessed using a Chi-squared test with a p-value cut off of <0.05. If the number of homozygous embryos was above the cut-off (i.e. in the expected 25% ratio), the allele was deemed to not cause a phenotype within the first 5 dpf. If the number of homozygous embryos was below the cut-off, the allele was carried forward into the second round of phenotyping. In the second round, we aimed to genotype 48 embryos for each phenotype, ideally from multiple clutches. An allele was documented as causing a phenotype if the phenotypic embryos were homozygous for the allele. We allowed up to 10% of embryos for a given phenotype to not be homozygous, to account for errors in egg collection. Such alleles were outcrossed for further genotyping with F4 embryos at a later date. Where possible, alleles were also submitted to complementation tests.
Kettleborough R.N., Busch-Nentwich E.M., Harvey S.A., Dooley C.M., de Bruijn E., van Eeden F., Sealy I., White R.J., Herd C., Nijman I.J., Fényes F., Mehroke S., Scahill C., Gibbons R., Wali N., Carruthers S., Hall A., Yen J., Cuppen E, & Stemple D.L. (2013). A systematic genome-wide analysis of zebrafish protein-coding gene function. Nature, 496(7446), 494-497.
Publication 2013
Adult Air Sacs Alleles Animals Blood Circulation Body Shape Brain Brain Death Cardiac Arrest Cell Death Cells Cranium Digestive System Eggs Embryo Epistropheus Fishes Genetic Complementation Test Genotype Heart Heterozygote Homozygote Hyperostosis, Diffuse Idiopathic Skeletal Intestines Liver Melanocyte Methanol Movement Muscle Tissue Mutagenesis Mutation Notochord Pancreas Phenotype Pigmentation Somites Touch Zebrafish
2
Calcium Sensor Imaging of Olfactory Responses in Drosophila
Flies were reared on standard cornmeal agar medium. We used the Gal4/UAS system (Brand et al., 1994 (link)) to direct the expression of the calcium sensors to PNs. GH146-Gal4 flies were a gift from L. Luo (Stanford University, Stanford, CA). All animals were adult females, 3–5 days after eclosion. Adult flies were dissected using previously described methods (Jayaraman and Laurent, 2007 ). Flies were anaesthetized in a vial on ice until movement stopped (<15 seconds) and then gently inserted into a hole in a piece of aluminum foil. Small drops of wax (55 °C) were used to suspend the fly in the hole, with the edge of foil defining a horizontal plane around the head and thorax, from the first antennal segment anteriorly to the scutellum posteriorly. The dorsal side of the foil was bathed in saline, while the ventral side (including antennae and maxillary palps) remained dry and accessible to odors. A window was cut in the dorsal head cuticle between the eyes, extending from the ocelli to the first antennal segment. Fat and air sacs dorsal and anterior to the brain were removed, but the perineural sheath was left intact. The proboscis was affixed with a small drop of wax to a strand of human hair to limit brain movement. Spontaneous leg movements were typically observed in this preparation for the duration of the recording (2–3 h). The saline composition used in all olfactory experiments was (in mM): 103 NaCl, 3 KCl, 5 N-tris (hydroxymethyl) methyl-2-aminoethane-sulfonic acid, 10 trehalose, 10 glucose, 26 NaHCO3, 1 NaH2PO4, 2.0 CaCl2, and 4 MgCl2, adjusted to 275 mOsm, pH 7.4.
Odors (different concentrations of octanol) were delivered using a custom-made odor-delivery system designed by Dmitry Rinberg, and a Teflon nozzle (entry diameter 1/8″) directed towards the antennae. Odors were delivered at different concentrations diluted in paraffin oil (Paraffin oil alone, 0.001%, 0.01%, 0.1%, 1.0% and 10%) in a constant stream of air (1 l/min) with an additional 10% dilution in air. For each concentration, five replicate deliveries were performed and the data averaged. Odor delivery times were measured using a mini-PID (Aurora Scientific Inc., Ontario, Canada). Odors were presented for 1s. All comparisons of sensor performance were made using experiments with identical odor presentation times. The results reported are based on data obtained from 5 GCaMP3-expressing flies (6 ALs) and 5 GCaMP5-expressing flies (6 ALs).
Akerboom J., Chen T.W., Wardill T.J., Tian L., Marvin J.S., Mutlu S., Calderón N.C., Esposti F., Borghuis B.G., Sun X.R., Gordus A., Orger M.B., Portugues R., Engert F., Macklin J.J., Filosa A., Aggarwal A., Kerr R.A., Takagi R., Kracun S., Shigetomi E., Khakh B.S., Baier H., Lagnado L., Wang S.S., Bargmann C.I., Kimmel B.E., Jayaraman V., Svoboda K., Kim D.S., Schreiter E.R, & Looger L.L. (2012). Optimization of a GCaMP Calcium Indicator for Neural Activity Imaging. The Journal of Neuroscience, 32(40), 13819-13840.
Publication 2012
Adult Agar Air Sacs Aluminum Animals Bicarbonate, Sodium Brain Calcium Chest Diptera DNA Replication Eye Glucose Hair Head Homo sapiens Magnesium Chloride Maxilla Movement Neoplasm Metastasis Obstetric Delivery Octanols Odors paraffin oils Saline Solution Sense of Smell Sodium Chloride Sulfonic Acids Technique, Dilution Teflon Trehalose Tromethamine Woman
3
Anesthesia and Dissection for Drosophila Imaging
Procedures for animal preparation were based on earlier methods (Turner et al., 2008 (link); Murthy and Turner, 2010 ). Briefly, flies were transferred to a glass tube and anesthetized on ice until movement ceased (about 15 s). A female fly was then gently inserted into a rectangular hole (about 0.77 by 1.5 mm) cut into a piece of aluminum foil glued to the underside of the recording platform. The fly’s head was tilted forward to provide access to the posterior surface of the brain where the KC cell bodies are located. The olfactory organs point downwards in this preparation, allowing airborne odor delivery (Figure 1A). The fly was fixed in place using fast-drying epoxy (Devcon 5-Minute Epoxy).
The bath surrounding the head capsule was continuously perfused with oxygenated saline (Wilson et al., 2004 (link)) and the cuticle at the back of the head was dissected away using sharpened forceps. We sometimes found it necessary to minimize brain motion by removing the pulsatile organ at the neck (care was taken to avoid damaging the gut) and the proboscis retractor muscles, which pass over the caudal aspect of the optic lobes. Air sacs and fat deposits occluding the MB were cleared from the brain’s surface. We did not purposefully attempt to remove the peri-neural sheath, as is needed for electrophysiological experiments. Flies remained healthy and active throughout the experiment, as evidenced by abundant voluntary leg movements. Many preparations were discarded due to excessive brain motion that prevented us from tracking individual neurons throughout the imaging session.
Honegger K.S., Campbell R.A, & Turner G.C. (2011). Cellular-Resolution Population Imaging Reveals Robust Sparse Coding in the Drosophila Mushroom Body. The Journal of Neuroscience, 31(33), 11772-11785.
Publication 2011
Air Sacs Aluminum Animals Bath Brain Capsule Cell Body Diptera Epoxy Resins Females Forceps Head MM 77 Movement Muscle Tissue Neck Nervousness Neurons Obstetric Delivery Odors Optic Lobe, Nonmammalian Saline Solution Sense of Smell
4
Zebrafish Behavioral and Developmental Assay
Fig 1 illustrates experimental perturbations (above the timeline), and associated analyses (below the timeline). Adult Tropical 5D zebrafish were housed at Sinnhuber Aquatic Research Laboratory at Oregon State University. All experiments were approved by the Institutional Animal Care and Use Committee (IACUC) of Oregon State University. Each tank was kept at 28°C on a 14h light/ 10h dark photoperiod. Group spawns of adult zebrafish were set up the night prior, and embryos were collected and staged [13 (link)]. Embryo chorions were enzymatically removed using pronase (90 μL of 25.3 U/μl; Roche, Indianapolis, In, USA) at 4 hours post fertilization (hpf) using a custom automated dechorionator and protocol described in Mandrell et al. [14 (link)]. Six hpf dechorinated embryos were placed individually into the wells of two 96-well plates per chemical using the automated embryo placement systems (AEPS) [14 (link)]. Chemicals were added to final well concentrations of 0, 0.0064, 0.064, 0.64, 6.4, and 64uM, with 0.64% DMSO included as the vehicle. Thus, there are 32 embryos per chemical per concentration. The layout of each concentration within a plate is shown in Fig 1. At 24hpf, an embryonic photomotor response (EPR) test was implemented [8 ]. After EPR, all exposed plates were wrapped with alumnium foil to prevent photodegradation, kept in a 28°C incubator, and statically exposed until 120hpf.
At 120hpf, zebrafish larvae movement was recorded in Viewpoint Zebrabox (Viewpoint Life Sciences, Lyon, France) during a 7-minute period of light followed by an 8-minute period of dark, then evaluated for 18 distinct morphological endpoints. The 18 morphological endpoints recorded for developmental assessment were Mortality (MORT), Yolk sac edema (YSE), Body axis (AXIS), Eye defect (EYE), Snout (SNOU), Jaw (JAW), Otic vesicle (OTIC), Pericardial edema (PE), Brain (BRAI), Somite (SOMI), Pectoral fin (PFIN), Caudal fin (CFIN), Pigmentation (PIG), Circulation (CIRC), Truncated body (TRUN), Swim bladder (SWIM), Notochord & Bent tail (NC), and Touch response (TR). Each morphological endpoint was recorded as a binary presence/absence according to the protocol detailed in [12 (link)]. All data were recorded by the Zebrafish Acquisition and Analysis Program (ZAAP) [12 (link)]. The current manuscript primarily focuses on the 120hpf behavioral assessment (see right-most ‘Light Condition Exposure’ portion of Fig 1).
Zhang G., Truong L., Tanguay R.L, & Reif D.M. (2017). A New Statistical Approach to Characterize Chemical-Elicited Behavioral Effects in High-Throughput Studies Using Zebrafish. PLoS ONE, 12(1), e0169408.
Publication 2017
Adult Air Sacs Brain Chorion Decompression Sickness Ear Edema Embryo Epistropheus Fertilization Human Body Hypomenorrhea Institutional Animal Care and Use Committees Larva Light Movement Notochord Pericardium Photodegradation Pigmentation Pronase Somites Sulfoxide, Dimethyl Tail TimeLine Touch Training Programs Yolk Sac Zebrafish
5
Chick Embryo Valproic Acid Injection
Freshly fertilized eggs of domestic chicks (Gallus gallus), of the Ross 308 (Aviagen) strain, were obtained from a local commercial hatchery (Agricola Berica, Montegalda (VI), Italy), placed in a cold room at 4 °C and maintained in a vertical position for 24–72 h. The eggs were then placed in the dark and incubated at 37.5 °C and 60% relative humidity, with rocking. The first day of incubation was considered embryonic day 0 (E0). Fertilized eggs were then selected by a light test on E14 before injection. Chick embryo injection was performed according to previous reports30 (link). Briefly, a small hole was made on the egg shell above the air sac, and 35 μmoles of VPA (Sodium Valproate, Sigma Aldrich) were administered to each fertilized egg, in a volume of 200 μl, by dropping the solution onto the chorioallantoic membrane. Age-matched control eggs were injected using the same procedure with 200 μL of vehicle (double distilled injectable water). After sealing the hole with paper tape, eggs were placed in a rocking incubator (FIEM srl, Italy) until E18, when eggs were placed in a hatching incubator (FIEM srl, Italy). Hatching took place at a temperature of 37.7 °C, with 60% humidity, as previously described16 (link). The day of hatching was considered post-hatching day 0 (P0). All subsequent procedures were performed in complete darkness, so that the chicks remained visually inexperienced until the moment of test. Two independent samples of chicks were used for the two experiments described.
Sgadò P., Rosa-Salva O., Versace E, & Vallortigara G. (2018). Embryonic Exposure to Valproic Acid Impairs Social Predispositions of Newly-Hatched Chicks. Scientific Reports, 8, 5919.
Publication 2018
Air Sacs Chick Embryo Chickens Cold Temperature Darkness Eggs Egg Shell Embryo Humidity Light Membrane, Chorioallantoic Sodium Valproate Strains Zygote

Most recents protocols related to «Air Sacs»

1
Automated Fish Swim Bladder Tracking
Algorithms were developed in C++ to determine the position of the center of mass of the swim bladder for each fish. A background image was initialized as the average of the first 2000 frames and subsequently updated at regular intervals during each session. For each frame, the image of each swim channel was processed in parallel in real time during image acquisition. The corresponding portion of the current background image was first subtracted from each channel image. The result was rectified, smoothed with a median 3x3 filter, eroded using a kernel size 2, pixel values scaled up to maximize contrast, and then thresholded to yield a binary image. Contours were extracted from the binary image with the border following algorithm of Suzuki and Abe [46 (link)] as implemented in OpenCV [47 ] and their centers of mass calculated and recorded for subsequent off-line analysis. As multiple contours were sometimes found in the same frame, a custom Python script using Markov chains was applied in a post-processing step to determine in this case which of the resulting candidate positions to accept as the most likely true position given the previous history.
Holman J.G., Lai W.W., Pichler P., Saska D., Lagnado L, & Buckley C.L. (2023). A behavioral and modeling study of control algorithms underlying the translational optomotor response in larval zebrafish with implications for neural circuit function. PLOS Computational Biology, 19(2), e1010924.
Publication 2023
Air Sacs Fishes Python Reading Frames
2
Acoustic Detection of Whale Prey Biomass
AZFP data were processed using Echoview (v.12; Echoview Software Pty Ltd.). Mean volume backscattering strength (Sv in dB re 1 m-1), a relative measure of density, was analyzed from 10 m below the surface to 5 m above the sounder-detected seafloor in bins of 5 m vertically by 5 pings horizontally. Sound speed [55 (link)] and absorption coefficients [56 (link)] were estimated at each frequency using temperature and salinity values reported for the closest oceanographic sampling station in Juan de Fuca Strait in September 2017 (approximately mid-Strait off Sooke Basin), as measured by Fisheries and Ocean Canada. Background noise was removed following the approach described in de Robertis & Higginbottom [57 (link)], using a minimum signal-to-noise ratio of 10 dB and maximum noise threshold of -125 dB re 1 m-1. Removal of acoustic noise was done through visual inspection of the echograms and applying filters following Ryan et al. [58 (link)]. Impulse and transient noise were removed with a maximum threshold of 10 dB and 12 dB, respectively. Backscatter in the processed echograms was scrutinized and classified based on echo morphology (shape and structure of the aggregations), depth distribution (including bottom association), and single target attributes. Mean volume backscattering strength (Sv) at 125 kHz was subtracted from that at 200 kHz to assess differences (MVBS200-125). Values greater or equal to 0 dB or lower or equal to 4 dB would be indicative of swimbladder-bearing fish, while MVBS 200–125 values greater than 5 dB would indicate backscatter dominated by zooplankton [59 (link)]. The processed volume scattering data were gridded into 1-min horizontal cells by 10-m vertical cells, then echo-integrated using an integration threshold of -70 dB over the whole water column into nautical area scattering coefficients (NASC; m2 nmi-2; [60 (link)]) to obtain relative measures of water column biomass where the whale was feeding. The overall pattern of log10-transformed NASC and the whale’s feeding rate during tag attachment were plotted in R, using a generalized additive model (GAM) with integrated smoothness estimation (y~s(x)). To estimate biomass per lunge count, we averaged NASC for each bottom-foraging time interval, and further averaged NASC across foraging intervals that had matching lunge counts to obtain a single estimate in each lunge-count category.
Reidy R., Gauthier S., Doniol-Valcroze T., Lemay M.A., Clemente-Carvalho R.B., Cowen L.L, & Juanes F. (2023). Integrating technologies provides insight into the subsurface foraging behaviour of a humpback whale (Megaptera novaeangliae) feeding on walleye pollock (Gadus chalcogrammus) in Juan de Fuca Strait, Canada. PLOS ONE, 18(3), e0282651.
Publication 2023
Acoustics Air Sacs Cells Cetacea ECHO protocol Fishes M Cells Multivesicular Body Salinity Sound Transients Zooplankton
3
Electrophysiological Recordings in Bengalese Finches
All procedures described below were approved by The Institutional Animal Care and Use Committee at Emory University. As described previously (Srivastava, Elemans, and Sober 2015 (link); Zia et al. 2020 (link); Zia et al. 2018 ), adult male Bengalese finches (>90 d old) were anesthetized using intramuscular injections of 40 mg/kg ketamine and 3 mg/kg midazolam injected and anesthesia was maintained using 1–5% isoflurane in oxygen gas. To record from the expiratory (respiratory) muscles, an incision was made dorsal to the leg attachment and rostral to the pubic bone and the electrode array was placed on the muscle surface using the “epimysial” approach described above. To record from syringeal (vocal) muscles, the vocal organ was accessed for electrode implantation via a midline incision into the intraclavicular air sac as described previously (Srivastava, Elemans, and Sober 2015 (link)) to provide access to the ventral syringeal (VS) muscle located on the ventral portion of the syrinx near the midline.
Chung B., Zia M., Thomas K.A., Michaels J.A., Jacob A., Pack A., Williams M.J., Nagapudi K., Teng L.H., Arrambide E., Ouellette L., Oey N., Gibbs R., Anschutz P., Lu J., Wu Y., Kashefi M., Oya T., Kersten R., Mosberger A.C., O’Connell S., Wang R., Marques H., Mendes A.R., Lenschow C., Kondakath G., Kim J.J., Olson W., Quinn K.N., Perkins P., Gatto G., Thanawalla A., Coltman S., Kim T., Smith T., Binder-Markey B., Zaback M., Thompson C.K., Giszter S., Person A., Goulding M., Azim E., Thakor N., O’Connor D., Trimmer B., Lima S.Q., Carey M.R., Pandarinath C., Costa R.M., Pruszynski J.A., Bakir M, & Sober S.J. (2023). Myomatrix arrays for high-definition muscle recording. bioRxiv.
Publication Preprint 2023
Adult Air Sacs Anesthesia Exhaling Finches Institutional Animal Care and Use Committees Intercostal Muscle Intramuscular Injection Isoflurane Ketamine Males Midazolam Muscle Tissue Ovum Implantation Oxygen Pubic Bone Syringomyelia Vocal Muscle
4
Surgical Implantation of Respiratory and Vocal Muscle Electrodes in Bengalese Finches
All procedures described below were approved by The Institutional Animal Care and Use Committee at Emory University. As described previously 20 (link),27 (link),42 (link), adult male Bengalese finches (>90 d old) were anesthetized using intramuscular injections of 40 mg/kg ketamine and 3 mg/kg midazolam injected and anesthesia was maintained using 1–5% isoflurane in oxygen gas. To record from the expiratory (respiratory) muscles, an incision was made dorsal to the leg attachment and rostral to the pubic bone and the electrode array was placed on the muscle surface using the “epimysial” approach described above. To record from syringeal (vocal) muscles, the vocal organ was accessed for electrode implantation via a midline incision into the intraclavicular air sac as described previously 20 (link) to provide access to the ventral syringeal (VS) muscle located on the ventral portion of the syrinx near the midline.
Chung B., Zia M., Thomas K.A., Michaels J.A., Jacob A., Pack A., Williams M.J., Nagapudi K., Teng L.H., Arrambide E., Ouellette L., Oey N., Gibbs R., Anschutz P., Lu J., Wu Y., Kashefi M., Oya T., Kersten R., Mosberger A.C., O’Connell S., Wang R., Marques H., Mendes A.R., Lenschow C., Kondakath G., Kim J.J., Olson W., Quinn K.N., Perkins P., Gatto G., Thanawalla A., Coltman S., Kim T., Smith T., Binder-Markey B., Zaback M., Thompson C.K., Giszter S., Person A., Goulding M., Azim E., Thakor N., O’Connor D., Trimmer B., Lima S.Q., Carey M.R., Pandarinath C., Costa R.M., Pruszynski J.A., Bakir M, & Sober S.J. (2023). Myomatrix arrays for high-definition muscle recording. bioRxiv.
Publication Preprint 2023
Adult Air Sacs Anesthesia Exhaling Finches Institutional Animal Care and Use Committees Intercostal Muscle Intramuscular Injection Isoflurane Ketamine Males Midazolam Muscle Tissue Ovum Implantation Oxygen Pubic Bone Syringomyelia Vocal Muscle
5
Evaluation of SifA-ELISA for Detecting Salmonella Infection in Chickens

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Publication 2023
Air Sacs Animals Antibody Formation Blood Chickens Cross Reactions Enzyme-Linked Immunosorbent Assay Escherichia coli Ethics Committees, Research Infection Intestines Liver Males Pasteurella multocida Salmonella Infections Serum Specific Pathogen Free Spleen

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More about "Air Sacs"

Avian Respiratory Structures: Exploring the Vital Role of Air Sacs in Avian Physiology and Biology Dive into the fascinating world of air sacs, the specialized respiratory structures found in birds and other avian species.
These thin-walled, balloon-like structures play a crucial role in the efficient delivery of oxygen to tissues and the removal of carbon dioxide, contributing to the unique breathing mechanism that enables powered flight in birds.
Unravel the intricate workings of air sacs, which connect to the lungs and facilitate the movement of air through the body, allowing for enhanced gas exchange during both inhalation and exhalation.
Gain insights into the structure and function of these respiratory marvels, and discover how they have evolved to adapt to the demands of avian physiology.
Delve into the latest research on air sacs, leveraging tools like MATLAB, MS-222, RNAlater, Digital cameras, TRIzol reagent, Tricaine, SZX16, Gryphax Arktur cameras, LM35JC10M sensors, and RNAlater solution to optimize your studies and uncover new discoveries.
Explore the broader implications of understanding avian respiratory systems, and how these findings can shape our understanding of avian biology and evolution.
Whether you're a researcher, a student, or simply fascinated by the natural world, this comprehensive overview of air sacs will provide you with a deeper appreciation for the intricate adaptations that enable birds to soar and thrive.
Embark on a journey of scientific exploration and unlock the secrets of these remarkable respiratory structures.