Ringraziamo Mike W. Davidson (Florida State University) per aver fornito il DNA plasmide codifica proteine ​​fluorescenti. Questo lavoro è stato sostenuto dal National Institutes of Health concedere GM72048 (a DWP).

1. Preparazione dei campioni goccia proteina fluorescente Una goccia proteina fluorescente del campione è costituito da una emulsione 1-octanol/water con la proteina fluorescente che risiedono nella fase acquosa. Questa emulsione è inserita tra un vetrino da microscopio e un quadrato 22 mm di vetro di copertura per le applicazioni di microscopia. <ol><li> Prima di effettuare campioni goccia proteina fluorescente il vetrini e vetri di copertura devono essere puliti e rivestiti con un agente idrofobo. </li><li> Pulire vetro lavando 5 minuti con acetone e lasciare asciugare per via aerea. (In alternativa, dopo aver pulito il vetro può essere trattato per 30 secondi e un detersivo plasma per ottenere risultati ottimali rivestimento). </li><li> Preparare una soluzione al 2% metiltrimetossisilano in acetone e rivestire il vetro durante una incubazione di 2 minuti in questa soluzione. Dopo il rivestimento rimuovere il vetro dalla soluzione e lasciare asciugare per via aerea. Poi risciacquare con etanolo al 70% da una bottiglia spray e lasciare asciugare di nuovo. (In alternativa, a questo punto il vetro può essere cotto al forno per 1 ora a 80 ° C per collegare covalentemente il rivestimento per il vetro). Vetro rivestito può essere conservato per almeno un mese. </li><li> Proteine ​​fluorescenti sono purificati come Suo proteine ​​6-tag da E. Coli 1. Misurare lo spettro di assorbimento delle proteine ​​purificate e preparare una diluizione stock con una densità ottica di circa 0,1 nel STE tampone (150 mM NaCl, 10 mM Tris-HCl pH 8, 1 mM EDTA), contenente 0,1% di albumina sierica bovina (BSA) . Inoltre preparare 10 ml di una miscela 1:1 di buffer di 1-ottanolo e STE in un tubo da 15 ml e mescolare energicamente. Dopo il congedo mescolando fino a quando la separazione di fase è completa. La fase superiore è l&#39;1-ottanolo. (Attenzione: poiché 1-ottanolo ha un odore forte, è importante usare un contenitore chiuso dei rifiuti per tutto ciò che viene a contatto con l&#39;1-ottanolo.) </li><li> Per rendere l&#39;emulsione pipetta 45 microlitri 1-ottanolo e 5 microlitri proteina fluorescente in una provetta per microcentrifuga. Toccare il tubo di un paio di volte con il dito per avviare la formazione di emulsione e poi sonicare il tubo per 30 secondi in un bagno di sonicazione. Nel frattempo ottiene un vetrino da microscopio rivestito e copertura in vetro pronto. Dopo sonicazione l&#39;emulsione deve essere completamente coperto. Immediatamente dopo la sonicazione pipetta 4 emulsione microlitri a partire dalla metà del tubo su un vetrino da microscopio ricoperto e coprire con un coperchio di vetro rivestito. </li><li> Se la procedura viene effettuata correttamente l&#39;emulsione deve diffondersi in modo uniforme tra il vetrino da microscopio e il vetro dell&#39;oggetto. In pochi minuti il ​​campione deve essere stabile, composto da circa 10 micron di spessore gocce fluorescenti con diametro variabile. La più grande goccioline sono vicino al centro del campione e le più piccole sono situate più verso i bordi. </li></ol> 2. Impostazione di un esperimento fotoconversione La procedura che segue è una strategia generale per la creazione di un esperimento fotoconversione proteina fluorescente. Questa procedura può essere applicata per le proteine ​​purificate così come per le cellule vive. <ol><li> I parametri che seguono forniscono un punto di partenza generale per impostare l&#39;esperimento fotoconversione: 40x 1.3NA olio obiettivo ad immersione Dimensione immagine = 512 x 512 pixel Scansione zoom = 4 Pixel tempo di permanenza = 6 msec. Z-risoluzione (dimensione del foro stenopeico) = 3 micron </li><li> Configurare due canali di rilevazione per la fluorescenza iniziale e photoconverted, così come un &quot;canale fotoconversione&quot;. In questo esempio useremo purificati mOrange2 proteina, che è un arancione-rosso photoconvertible proteina fluorescente. La specie arancione viene rilevato utilizzando 561 nm di eccitazione e la fluorescenza viene raccolto tra 570 nm e 630 nm. La specie photoconverted rosso viene rilevato utilizzando 633 nm di eccitazione e la fluorescenza viene raccolto tra 640 nm e 700 nm. Per il &quot;canale fotoconversione&quot; seleziona 488 nm di eccitazione e raccogliere la fluorescenza tra 490 nm e 540 nm. (Nota:. Immagini del canale fotoconversione non è strettamente necessario) </li><li> Utilizzare il canale per l&#39;imaging di fluorescenza iniziale con scansione continua per regolare la potenza del laser e rivelatore di guadagno per una qualità ottimale delle immagini. </li><li> Attivare il canale fotoconversione e selezionare un laser a bassa potenza. Avviare una serie di immagini lasso di tempo e di aumentare gradualmente il laser fotoconversione fino significativo sbiancamento della fluorescenza iniziale si osserva. Continuare la scansione fino a quando la fluorescenza iniziale è circa il 75% sbiancato. </li><li> Disattivare il canale fotoconversione e attivare il canale di rilevazione per la fluorescenza photoconverted. Avvio di imaging con un guadagno del rivelatore alta e potenza del laser a bassa e aumentare gradualmente la potenza del laser fino a quando la fluorescenza photoconverted viene rilevato. Una volta rilevare la fluorescenza photoconverted è possibile regolare la potenza del laser e rivelatore di guadagno per una qualità ottimale delle immagini. </li><li> Infine, la potenza del laser utilizzatoper fotoconversione così come la durata della fotoconversione bisogno di essere ottimizzato. Aumentare la potenza del laser fotoconversione accelererà la velocità di fotoconversione, potenza del laser però troppa photobleach la proteina. </li><li> Una volta che la potenza del laser fotoconversione ottimale e la durata sono stati determinati, questi parametri possono essere utilizzati per configurare una photobleaching standard o modulo FRAP e il &quot;canale fotoconversione&quot; non è più necessaria. </li></ol> 3. Evidenziando ottico a doppia sonda con mOrange2 e Dronpa A causa del rosso-spostato proprietà spettrali, mOrange2 può essere utilizzato in combinazione con il verde Dronpa photoswitchable proteina fluorescente per evidenziare ottici a doppia sonda per consentire l&#39;evidenziazione selettiva di 4 celle singole (organello) popolazioni. <ol><li> Le cellule sono coltivate in piatti con fondo trasparente MatTek e transfettate 24 ore prima immagine che utilizzano standard di trasfezione Lipofectamine2000 1. </li><li> Impostare il microscopio per mOrange2 fotoconversione come descritto nella sezione 2. </li><li> Configurare il microscopio per photoswitching Dronpa. Dronpa fluorescenza può essere ripreso con il mOrange2 &quot;canale fotoconversione&quot; (vedi punto 2.2). (Nota: Ridurre al minimo la potenza del laser utilizzato per l&#39;imaging Dronpa, perché troppa potenza del laser provoca l&#39;inattivazione di Dronpa.) Aggiungere un canale per fotoattivazione Dronpa. Utilizziamo 800 nm a due fotoni di eccitazione per fotoattivazione, ma in alternativa questo può essere realizzato utilizzando 405 nm di eccitazione. Determinare la potenza del laser necessaria per l&#39;imaging, fotoattivazione e photoinactivation di Dronpa fluorescenza. </li><li> Attenzione: fotoconversione di mOrange2 e inattivazione di Dronpa sia insorgere a seguito di eccitazione 488 nm. A causa della potenza del laser ad alta richiesta per mOrange2 fotoconversione questo sarà anche inattivare Dronpa fluorescenza. D&#39;altra parte, l&#39;inattivazione Dronpa avviene già in potenza del laser molto più basso e può essere eseguito senza significativi fotoconversione mOrange2. </li><li> Una volta che i parametri per fotoconversione mOrange2 e photoswitching Dronpa sono impostati, evidenziando sonda a doppia ottica si realizza attraverso i seguenti passaggi. In primo luogo, inattivare Dronpa fluorescenza in tutto il campo di vista con bassa potenza di eccitazione 488 nm. In secondo luogo, selezionare una regione di interesse e mOrange2 photoconvert ad alto potere di eccitazione 488 nm. Infine, selezionare una regione di interesse per attivare Dronpa fluorescenza. </li></ol> 4. Rappresentante Risultati <img src="/files/ftp_upload/1995/1995fig1.jpg" alt="Figura 1" /> Figura 1. Droplet preparazione del campione. A) campione goccia correttamente preparati. B) del campione preparato senza rivestimento del vetrino da microscopio e vetro di copertura. C) del campione preparato senza l&#39;aggiunta di 0,1% BSA. <img src="/files/ftp_upload/1995/1995fig2.jpg" alt="Figura 2" /> Figura 2. Effetto della potenza fotoconversione laser e la durata in mOrange2 fotoconversione. Goccioline contenenti mOrange2 singola proteina sono stati continuamente photoconverted utilizzando diverse quantità di potenza di 488 nm laser. Potenza del laser utilizzato per fotoconversione relativa era del 10% (solido), il 25% (tratteggiata), e 100% (tratteggiata). A) specie arancio fluorescente. B) Photoconverted specie rosso fluorescente. <img src="/files/ftp_upload/1995/1995fig3.jpg" alt="Figura 3" /> Figura 3. Evidenziando ottico a doppia sonda con mOrange2 e Dronpa. A) la cellula che esprime mOrange2-istoni H2B e Dronpa-Mito prima fotoconversione, mostrando fluorescenza arancione nel nucleo e fluorescenza verde nei mitocondri. B) Dronpa fluorescenza è stato spento con bassa potenza di eccitazione 488 nm, causando fotoconversione minimo di mOrange2. C) mOrange2 era photoconverted al rosso ad alto potere di eccitazione 488 nm. D) Dronpa fluorescenza è stata riattivata con 800 nm 2-fotone di eccitazione. I pannelli sono sovrapposizioni delle immagini a fluorescenza con l&#39;immagine di contrasto differenziale di interferenza.

<ol>
	<li>Kremers, G. J., Hazelwood, K. L., Murphy, C. S., Davidson, M. W., Piston, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photoconversion+in+orange+and+red+fluorescent+proteins.">Photoconversion in orange and red fluorescent proteins.</a> Nature Methods. 6, 355-358 (2009).</li></ol>

Nessun conflitto di interessi dichiarati.

Il campione purificato goccia proteina fluorescente è un formato di campione molto comodo per la caratterizzazione fotofisiche di proteine ​​fluorescenti, ad esempio per studiare photobleaching cinetica e cinetica fotoconversione. Il volume delle gocce estremamente piccole (~ 20 picolitri) facilita photobleaching ed esperimenti fotoconversione, che può essere difficile da eseguire in sistemi basati cuvetta. Inoltre, come mostrato qui il campione goccia è ideale per la creazione di un microscopio confocale per le ...

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<td>Microsope slides</td>
<td>&nbsp;</td>
<td>VWR Scientific</td>
<td>48312-003</td>
<td>&nbsp;</td>
<tr>
<td>22 mm cover glass</td>
<td>&nbsp;</td>
<td>Corning</td>
<td>2940-245</td>
<td>&nbsp;</td>
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<td>1-octanol</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>O4500</td>
<td>&nbsp;</td>
<tr>
<td>methyltrimethoxysilane</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>M6420</td>
<td>&nbsp;</td>
<tr>
<td>MatTek dishes</td>
<td>&nbsp;</td>
<td>MatTek Corp.</td>
<td>P35G-1.5-14-C</td>
<td>&nbsp;</td>
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<td>Lipofectamine2000</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
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photoconversion of purified fluorescent proteins and dual-probe optical highlighting in live cells

Photoconvertible proteine ​​fluorescenti (pc-PQ) sono una classe di proteine ​​fluorescenti con &quot;ottica evidenziatore&quot; capacità, il che significa che il colore della fluorescenza può essere cambiato l&#39;esposizione alla luce di una specifica lunghezza d&#39;onda. Evidenziando ottico permette la marcatura non invasiva di una sottopopolazione di molecole fluorescenti, ed è quindi ideale per il monitoraggio singole cellule o organuli. Parametri critici per fotoconversione efficiente sono l&#39;intensità e il tempo di esposizione della luce fotoconversione. Se l&#39;intensità è troppo bassa, fotoconversione sarà lenta o non verificarsi affatto. D&#39;altra parte, l&#39;intensità troppo o troppo a lungo possibile l&#39;esposizione photobleach la proteina e quindi ridurre l&#39;efficienza della fotoconversione. Questo protocollo descrive un approccio generale come impostare un microscopio confocale a scansione laser per pc-FP applicazioni fotoconversione. In primo luogo, si descrive una procedura per la preparazione di campioni di gocce di proteine ​​purificate. Questo formato campione è molto conveniente per lo studio del comportamento fotofisiche di proteine ​​fluorescenti al microscopio. In secondo luogo, useremo il campione goccia proteina per mostrare come configurare il microscopio per fotoconversione. E, infine, mostreremo come di evidenziare la ottici in cellule vive, inclusa l&#39;evidenziazione ottica dual-sonda con mOrange2 e Dronpa.

Watch this Scientific Journal Video about Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells at JoVE.com

Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Questo protocollo descrive un approccio generale per eseguire fotoconversione di proteine ​​fluorescenti su un microscopio confocale a scansione laser. Si descrivono le modalità per la fotoconversione di campioni di proteine ​​puried, nonché per evidenziare ottici a doppia sonda in cellule vive con mOrange2 e Dronpa.

Fotoconversione di proteine ​​purificate fluorescenti ed evidenziazione ottica dual-sonda in cellule vive

Photoconvertible fluorescent proteins (pc-FPs) are a class of fluorescent proteins with "optical highlighter" capability, meaning that the color of ...

photoconversion-purified-fluorescent-proteins-dual-probe-optical

Research

JoVE Journal

Biology

12.1K Views.  Vanderbilt University. Questo protocollo descrive un approccio generale per eseguire fotoconversione di proteine ​​fluorescenti su un microscopio confocale a scansione laser. Si descrivono le modalità per la fotoconversione di campioni di proteine ​​puried, nonché per evidenziare ottici a doppia sonda in cellule vive con mOrange2 e Dronpa.

Video: Photoconversion of Purified Fluorescent Proteins and Dual-probe Optical Highlighting in Live Cells

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Etichettatura Cellule staminali con le tinture fluorescenti per la rilevazione non invasiva con Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

Ricerca

Biologia

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Manutenzione e pulizia adeguata del microscopio

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

Etichettatura fluorescente di COS-7 Esprimendo SNAP-tag proteine ​​di fusione per l&#39;imaging cellulare dal vivo

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic development 1, 2. One of the signaling pathways involved in this regulation is the Wnt pathway, which is used many times during embryogenesis and critical for human disease3, 4, 5. Multiple molecular components of this pathway coordinately regulate signaling in a spatially-restricted manner, but the underlying mechanisms are not fully understood. Xenopus embryonic epithelial cells is an excellent system to study subcellular localization of various signaling proteins. Fluorescent fusion proteins are expressed in Xenopus embryos by RNA microinjection, ectodermal explants are prepared and protein localization is evaluated by epifluorescence. In this experimental protocol we describe how subcellular localization of Diversin, a cytoplasmic protein that has been implicated in signaling and cell polarity determination6, 7 is visualized in Xenopus ectodermal cells to study Wnt signal transduction8. Coexpression of a Wnt ligand or a Frizzled receptor alters the distribution of Diversin fused with red fluorescent protein, RFP, and recruits it to the cell membrane in a polarized fashion 8, 9. This ex vivo protocol should be a useful addition to in vitro studies of cultured mammalian cells, in which spatial control of signaling differs from that of the intact tissue and is much more difficult to analyze.

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus embryonic ectoderm has become an attractive model for studies of cell polarity. An assay is described, in which subcellular distribution of fluorescent proteins is assessed in ectoderm cells. This protocol will help address questions related to spatial control of signaling.

Traslocazione polarizzato di proteine ​​fluorescenti in Xenopus</emEctoderma> in risposta alla segnalazione Wnt

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

Vivo Imaging delle proteine ​​GFP-etichettati in<em&gt; Drosophila</em&gt; Gli oociti

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

4D Imaging di aggregazione proteica in cellule vive

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Intravitale Microscopia per le strutture Imaging subcellulari in topi vivi Esprimere proteine ​​fluorescenti

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

In vivo clonale Monitoraggio di staminali ematopoietiche e cellule progenitrici Contrassegnato da cinque proteine ​​fluorescenti che utilizzano confocale e microscopia Multiphoton

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Functional Reconstitution and Channel Activity Measurements of Purified Wildtype and Mutant CFTR Protein

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of transporters. The phosphorylation and nucleotide dependent chloride channel activity of CFTR has been frequently studied in whole cell systems and as single channels in excised membrane patches. Many Cystic Fibrosis-causing mutations have been shown to alter this activity. While a small number of purification protocols have been published, a fast reconstitution method that retains channel activity and a suitable method for studying population channel activity in a purified system have been lacking. Here rapid methods are described for purification and functional reconstitution of the full-length CFTR protein into proteoliposomes of defined lipid composition that retains activity as a regulated halide channel. This reconstitution method together with a novel flux-based assay of channel activity is a suitable system for studying the population channel properties of wild type CFTR and the disease-causing mutants F508del- and G551D-CFTR. Specifically, the method has utility in studying the direct effects of phosphorylation, nucleotides and small molecules such as potentiators and inhibitors on CFTR channel activity. The methods are also amenable to the study of other membrane channels/transporters for anionic substrates.

Described here is a rapid and effective procedure for functional reconstitution of purified wild-type and mutant CFTR protein that preserves activity for this chloride channel, which is defective in Cystic Fibrosis. Iodide efflux from reconstituted proteoliposomes mediated by CFTR allows studies of channel activity and the effects of small molecules.

Ricostituzione funzionale e attività Canale Misure di purificata di tipo selvatico e mutante CFTR proteine

The Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) is a unique channel-forming member of the ATP Binding Cassette (ABC) superfamily of ...

Characterization of Calcification Events Using Live Optical and Electron Microscopy Techniques in a Marine Tubeworm

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH distribution and calcium ions can be observed using live microscopy over time. This allows identification of the life stage and the tissue with the feature of interest for further electron microscopy studies. Life stage and tissues of interest are typically higher in pH and Ca signals. 
Here, using H. elegans, we present a protocol to characterize the presence of calcium carbonate structures in a biological specimen on the scanning electron microscope (SEM), using energy-dispersive X-ray spectroscopy (EDS) to visualize elemental composition, using electron backscatter diffraction (EBSD) to determine the presence of crystalline structures, and using transmission electron microscopy (TEM) to analyze the composition and structure of the material. In this protocol, a focused ion beam (FIB) is used to isolate samples with dimension suitable for TEM analysis. As FIB is a site specific technique, we demonstrate how information from the previous techniques can be used to identify the region of interest, where Ca signals are highest.

We demonstrate the use of various microscopy methods that are useful in observing the calcification of a tubeworm, Hydroides elegans, as well as locating and characterizing the first calcified material. Live microscopy and electron microscopy are used together to provide functional and material information that are important in studying biomineralization.

Caratterizzazione di calcificazione eventi utilizzando dal vivo ottica ed elettronica Microscopia tecniche in un tubeworm Marine

Characterizing the first event of biological production of calcium carbonate requires a combination of microscopy approaches. First, intracellular pH ...

Fotoconversione di proteine ​​purificate fluorescenti ed evidenziazione ottica dual-sonda in cellule vive

Fotoconversione di proteine purificate fluorescenti ed evidenziazione ottica dual-sonda in cellule vive