This work was supported in part by NIH grants (1R01CA137494, R01CA132115, R01CA086072 to R.G.P.), the Kimmel Cancer Center NIH Cancer Center Core grant P30CA056036 (R.G.P.), a grant from the Breast Cancer Research Foundation, generous grants from the Dr. Ralph and Marian C. Falk Medical Research Trust (R.G.P.) and a grant from the Pennsylvania Department of Health (R.G.P.). In part this work was supported by an American Italian Cancer Foundation postdoctoral fellowship (G.D.) and Bioimaging Shared Resource of the Sidney Kimmel Cancer Center (NCI 5 P30 CA-56036).The Department specifically disclaims responsibility for an analysis, interpretations or conclusions. There are no conflicts of interest associated with this manuscript.

Hem ifade vektörleri EYFP-Parkin ve pDsRed2-Mito ile fibroblast 1. elektroporasyonu <ol><li> 100 mg / ml DMEM (Dulbecco değiştirilmiş Eagle ortamı),% 10 fetal bovin serumu, 2 mmol / L L-glutamin, 100 U / ml penisilin ortamı kullanılarak 10 sm doku kültürü plakasında ölümsüzleştirilmiş fare embriyonik fibroblast hücreleri büyür ve 37 ° C&#39;de,% 5 CO2 içeren nemli bir atmosferde kültürlendi. <ol><li> % 80 hücre Akışların birleştiği yerde hücreler, HPO 4, KH 2.4 g, steril emilerek tam bir DMEM ortamıyla atmak ve steril 1 x fosfat tampon çözeltisi (10 ml PBS) (NaCl 80 g, KCl 2.0 g, Na 2, 14.4 g ekleme 7.4): 2 PO 4 ve 1 L H2O PH damıtıldı. </li><li> Steril emme PBS atın ve Tripsin-EDTA% 0.25 1 ml ekleyin. (- 3 dk 2) Hücreler müstakil kadar 37 ° C&#39;de inkübe plakası. _, Tam DMEM orta 4 ml ekleyin hücreleri tekrar süspansiyon ve 10 çıkar6; hemositometre sayımı hücre süspansiyonu l. Not: 16 Köşe kareler bir takım hücre sayısı hücrelerin x 10 4 / mi sayısına eşit olacak şekilde hemositometre tasarlanmıştır. </li></ol></li><li> Elektroporasyon işlemine 24 saat önce 10 cm doku kültürü yemekleri üzerine Tohum 1 x 10 6 hücre. </li><li> Steril emilerek tam bir DMEM ortamıyla atın ve steril 10 ml PBS ilave edin. Steril emme PBS atın ve Tripsin-EDTA% 0.25 1 ml ekleyin. (- 3 dk 2) Hücreler müstakil kadar 37 ° C&#39;de inkübe plakası. tam bir DMEM ortamıyla 4 ml ilave edilir ve 15 ml&#39;lik bir tüp içinde tekrar süspansiyon hücreleri. </li><li> bir buzdolabında-santrifüj (4 ° C) ile 5 dakika boyunca 250 xg&#39;de hücreleri aşağı doğru döndürün. Steril emme Süpernatantı atın ve steril PBS 1 ml pelletini. 4 ° C&#39;de 5 dakika boyunca 250 xg&#39;de hücreleri aşağı doğru döndürün. </li><li> Steril emerek supernatant atmak ve 100 ul o ekleyinelektroporasyon f çözüm karışımı hücre pelet (82 Company solüsyon 1 elektroporasyon Çözüm V + 18 ul ul) ve hafifçe pipetleme pelletini (Daha fazla bilgi için kullanım kılavuzuna başvurun). hücre süspansiyonu de EYFP-Parkin (uyarma / emisyon 514/527) ve 1 mcg pDsRed2-Mito (uyarma / emisyon 565/620) 2 ug ekleyin. </li><li> tek kullanımlık pastör kullanarak steril küvete solüsyonu (her iki araç kiti ile birlikte sağlanır) aktarın ve önceden ayarlanmış bir program NIH / 3T3 U-030 (Tek nabız, gerilim 200 V, kapasitans 960 mF, darbe süresi 20 milisaniye kullanarak electroporate , darbe sayısı: 1). </li><li> 37 ° C&#39;de,% 5 CO2 içeren nemli bir atmosferde bunları hemen Elektroporasyon işleminden sonra, taze, önceden ısıtılmış tam bir DMEM ortamıyla 500 ul ve 6 cm doku kültürü plakası, canlı görüntü dereceli hücreyi tohum ve inkübe 24 saat. </li></ol> 2. Time-Lapse video Mikroskopi <ol><li&gt; Kullanmadan önce 37 ° C&#39;de mikroskop odasının sıcaklığı ayarlayın. </li><li> Bu noktada deney protokolü ile devam etmek için, belirli bir canlı görüntüleme ortamı hazırlar. aşağıdaki gibi canlı görüntüleme ortamı hazırlayın; karıştırın DMEM fenol içermeyen,% 10 fetal bovin serumu, 2 mmol / L L-glutamin, 100 U / ml penisilin ve 100 mg / ml streptomisin ile takviye edilmiştir. Bir su banyosu içinde 37 ° C &#39;de, canlı görüntü ortamı önceden ısıtın. </li><li> Steril emilerek elektroporasyona hücre ortamı atılır ve P1000 pipet kullanarak, önceden ısıtılmış canlı görüntüleme ortamının 1 ml ilave edilir ve 37 ° C&#39;de 30 dakika süreyle plaka inkübe edin. </li><li> 37 ° C&#39;lik bir sabit sıcaklıkta önce mikroskop odasındaki Plaka, inkübasyon süresi, akış,% 5 CO2 içinde. </li><li> büyük hareketler ya da salınımları kaçınarak mikroskobun odasına Electroporated hücreleri ile plaka koyun. </li><li> Yazılım arayüzü kullanarak, her iki fluore tespit etmek için mikroskop ayarlamakko-transfekte vektörler tarafından kodlanan füzyon proteinleri scence sinyalleri EYFP-Parkin ve (Uyarım aralığı 495 nm ve emisyon aralığı yeşil 520-550 nm, 510) pDsRed2-Mito (Uyarım maksimum 558 nm ve emisyon maksimum 583 nm, kırmızı). <ol><li> time-lapse video mikroskopi yazılımını açın. Üst menüde pDsRed2-Mito için EYFP-Parkin ve rodamin için FITC seçin. Üst menüde büyütme (20X) seçin. </li></ol></li><li> yazılım arayüzü kullanarak, hem birlikte transfekte vektörleri ifade tek bir hücre için bakmak ve konumunu kaydeder. 10 hücrelerinin en az her deneysel koşul (Deney grubu) için toplanana kadar bu adımı tekrarlayın. <ol><li> menu &quot;Uygulamalar&quot; ı seçin ve Çok Boyutlu Edinme tıklayın. Çok Boyutlu Toplama pencerelerinde bu tür satın almalar sayısı, her satın alma ve kaydedilen hücrenin konumu arasındaki zaman aralığı olarak gerekli parametreleri seçin. Edinme &quot;üzerine tıklayın&quot;Satın alma işlemini başlatmak için. </li></ol></li><li> EYFP-Parkin ve 15 dakika olmak üzere toplam aralığı için görüntüleri her 5 dakikada toplama DsRed2-Mito floresan sinyal hem bazal alımı başlatabilirsiniz. </li><li> son çalışma konsantrasyonu iki katından daha yüksek bir FCCP bir konsantrasyon ile önceden ısıtılmış canlı görüntüleme ortamı hazırlayın (0,1-10 uM, hücre tipine bağlı olarak). </li><li> satın alma işlemini kesintiye ve P1000 pipet kullanarak mikroskobun odasının içine plakasına FCCP önceden ısıtılmış canlı görüntüleme ortamı yavaşça 1 ml ekleyin. </li><li> daha önce kaydedilmiş konumu kullanarak, 3 saatlik bir toplam süre için, tarif edildiği gibi elde etme işlemi başlatın. bunları analiz ve satın alma bittiğinde mitophagic sürecinin bir video oluşturmak için tüm kazanılmış görüntüleri kaydedin. </li></ol> 3. Analiz <ol><li> Bir kişisel bilgisayarda &quot;.tiff&quot; biçiminde tüm kazanılmış görüntüleri toplayın </li><li> Bir grafik yumuşak görüntüleri açıngereçleri ve zamansal aralık define (Görüntüler her 5 dakika edinilen) mitokondrial membran içine Parkin istihdamı gösteren ilk görüntüye FCCP mitokondriyal kaynaklı-depolarizasyon dan oluştu. Deney grubunda, her bir hücre için, bu analiz tekrarı. </li><li> Deney grubu adıyla bir tablo üzerinde sütunun ilk hücresini Etiket. her bir deney grubu için, 10 hücreleri en az Parkin alımı için zamansal aralıkları ölçmek. Parkin işe alım için hesaplanan geçici aralıklarla ortalama yapın ve her bir deney grubu için standart sapmayı tanımlamak (Ortalama ± SD, n = 10). <ol><li> sütun tek hesaplanan geçici aralıklarla içine düzenleyin. Her sütun n = deney grubunun 10 ölçümlerini içerir. </li><li> Formula Builder menüden fonksiyon &quot;ortalama&quot; seçeneğini seçin. sütunundaki verileri seçin. Formula Builder menüden fonksiyon &quot;Standart Sapma&quot; seçeneğini seçin. Selesütundaki veriler, CT. </li></ol></li><li> istatistiksel bir yaklaşım kullanarak, deney grupları arasında mitophagy dinamiği önemli bir fark tanımlamak için uygun istatistik uygulayarak deney grupları karşılaştırmak. (Örneğin, iki grup = Student t-testi, üç veya daha fazla gruplar = ANOVA artı bir post-hoc testi) </li></ol>

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The authors have nothing to disclose.

Time-lapse mikroskopi uzun süre boyunca hücresel dinamiklerinin gözlem zamanında tek bir gözlem canlı hücre görüntüleme uzanan tekniği olarak tanımlanabilir. bu gözlemci gerçek zamanlı olarak tek bir floresan etiketli protein tespit ve tek bir canlı hücrenin içinde kendi dinamiğini takip sağlar, çünkü bu yöntem, basit bir konfokal veya canlı hücre mikroskopi ayrılır. Aslında, konfokal mikroskopi kolayca flüoresan antikor kullanılarak imüno-etiketli proteinlerin tanımlar, ancak, hücre ve...

Macro autophagy is an intracellular process that involves the catabolic degradation of both damaged and dysfunctional cellular components, such as organelles and proteins for the purpose of either recycling or energy production. To initiate this metabolic process, the cell engulfs the damaged cellular components into a double-membrane structure, known as an autophagosome, which fuses with a lysosome and its content is degraded and recycled 1,2. There are two major types of autophagy, the non-selective and selective. The non-selective autophagy process occurs when the cell is under nutrient deprivation conditions and needs to scavenge for both essential nutrients and energy. However, selective autophagy occurs to mediate the removal of both dysfunctional/damaged organelles and proteins that otherwise could be toxic. One of the most studied selective autophagy process is the removal of mitochondria, termed mitophagy 1,3-5.
Mitochondria are the central organelles for cell metabolism and the primary source of adenosine triphosphate (ATP) via oxidative phosphorylation through the electron transport chain, fatty acid oxidation, and tricarboxylic acid (TCA) cycle. Moreover, mitochondria regulate reactive oxygen species (ROS) production and release proteins that participate in cell death pathways 6-8.
PTEN-induced putative kinase 1 (PINK1) and Parkin RBR E3 ubiquitin ligase (Parkin) are the key proteins implicated in the mitophagy process. Parkin can protect against cell death by keeping the cell healthy through mitochondrial quality control9. Upon the loss of mitochondrial membrane potential, cytosolic Parkin is recruited to the mitochondria by PINK1. This recruitment triggers the sequential events of mitophagy 10. There is a broad range of evidence that mitophagy is a fundamental mitochondria quality control process and abnormalities in this process drive disease 7. For instance, autosomal recessive Parkinson's disease has been associated with mutations in the genes that encode for Parkin and PINK1 (PARK2 and PINK1, respectively) 11. The quality control of mitochondrial health is essential for the removal of mitochondria that contribute to the accumulation of ROS12. Excessive presence of intracellular ROS can lead to damage of both nuclear and mitochondrial DNA (DNA and mt DNA, respectively).
Herein, we show a time-lapse video microscopy approach to follow the aggregation of Parkin after the induction of Parkin-mediated mitophagy in immortalized mouse embryonic fibroblasts via in vitro administration of carbonyl cyanide 4-(trifluoromethoxy)-phenylhydrazone (FCCP), an uncoupling agent. FCCP disrupts ATP synthesis by short circuiting protons across the outer mitochondria membrane and hence uncoupling oxidative phosphorylation from the electron transport chain 13. Triggering the depolarization of the mitochondrial membrane leads to the disruption of mitochondria and selective Parkin-dependent removal. Therefore, transfecting the cells of interest with an expression vector encoding Parkin fused with a fluorescent marker (enhanced yellow fluorescent protein, EYFP) can be used as a fluorescent tag to follow the recruitment of Parkin during the mitophagic process. In order to visualize the mitochondria, we co-transfected pDsRed2-Mito, which encodes red fluorescent protein (DsRed2) that contains a mitochondrial targeting sequence of cytochrome c oxidase subunit VIII (Mito). pDsRed2-Mito is designed for fluorescent labeling of mitochondria14. The time required for Parkin translocation into the mitochondrial membrane can be measured and gives an indirect measure of cellular health. For example, we can say that if a cell line knocked-out for a particular gene of interest shows either a faster or slower recruitment of Parkin after the induction of mitophagy by FCCP, that gene product would be a key player in order to keep the metabolic rates of the cell at the physiological status and prevent the development of diseases. Therefore, the time-lapse video microscopy provides a very powerful tool for both basic and clinical research applications in following the dynamic of labeled proteins during their molecular processes and understanding how these processes are affected during a pathological condition.

<table><tbody><tr><td>DMEM</td><td>Corning Life Science</td><td>10-013-CV</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>Phenol-free DMEM</td><td>Corning Life Science</td><td>17-205-CV</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>Fetal Bovine Serum</td><td>Sigma-Aldrich</td><td>F2442</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>L-Glutamine</td><td>Gibco</td><td>25030</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>Penicillin/streptomycin</td><td>Corning Life Science</td><td>30-002-CI</td><td>Pre-warm at 37 &amp;#176;C before use</td></tr><tr><td>EYFP-PARKIN expression vector</td><td>Addgene</td><td>23955</td><td></td></tr><tr><td>pDsRed2-Mito expression vector</td><td>Clontech</td><td>632421</td><td></td></tr><tr><td>Nucleofector 2B device</td><td>LONZA</td><td>AAD-1001S</td><td></td></tr><tr><td>Nucleofector for kit R NIH/3T3</td><td>LONZA</td><td>VCA-1001</td><td></td></tr><tr><td>ZEISS AXIOVERT 200M inverted microscope</td><td>CARL ZEISS</td><td></td><td></td></tr><tr><td>Carbonyl Cyanide 4-(trifluoromethoxy)-Phenylhydrazone (FCCP)</td><td>Sigma-Aldrich</td><td>C2920</td><td></td></tr><tr><td>MetaMorph</td><td>Molecular Devices</td><td>Experimental Builder</td><td></td></tr><tr><td>ImageJ</td><td>National Institute of Health</td><td>Experimental Builder</td><td></td></tr></tbody></table>

time-lapse video microscopy for assessment of eyfp-parkin aggregation as a marker for cellular mitophagy

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time points. Time intervals can be set through a computer interface that controls the microscope-integrated camera. This kind of microscopy requires both the ability to acquire very rapid events and the signal generated by the observed cellular structure during these events. After the images have been collected, a movie of the entire experiment is assembled to show the dynamic of the molecular events of interest. Time-lapse video microscopy has a broad range of applications in the biomedical research field and is a powerful and unique tool for following the dynamics of the cellular events in real time. Through this technique, we can assess cellular events such as migration, division, signal transduction, growth, and death. Moreover, using fluorescent molecular probes we are able to mark specific molecules, such as DNA, RNA or proteins and follow them through their molecular pathways and functions. Time-lapse video microscopy has multiple advantages, the major one being the ability to collect data at the single-cell level, that make it a unique technology for investigation in the field of cell biology. However, time-lapse video microscopy has limitations that can interfere with the acquisition of high quality images. Images can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. Herein, we describe a protocol for the dynamic acquisition of a specific protein, Parkin, fused with the enhanced yellow fluorescent protein (EYFP) in order to track the selective removal of damaged mitochondria, using a time-lapse video microscopy approach.

Bu yazıda, time-lapse video mikroskopi tek bir hücrede floresan etiketli proteinlerin moleküler olayları takip etmek için kullanılabilecek güçlü bir tekniktir nasıl gösterir. temsilcisi sonuçları da bu tekniğin yüksek kaliteli görüntüler elde edilmesi için izin verir nasıl gösterir. Moleküler sürecinin görüntüleri elde edilir, biz farklı şekillerde bunları analiz için fırsat var. Burada, biz seçici hasarlı mitokondri kaldırarak hücresel homeostazı koru...

Watch this Scientific Journal Video about Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy at JoVE.com

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

Hücresel Mitophagy için bir Marker olarak EYFP-Parkin Toplama Değerlendirme Time-Lapse video Mikroskopi

Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time ...

time-lapse-video-microscopy-for-assessment-eyfp-parkin-aggregation-as

Research

JoVE Journal

Biology

7.1K Görüntüleme.  Thomas Jefferson University, SKCC. Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Video: Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a quality control mechanism that can target and selectively degrade cellular material including organelles1-4. For example, damaged or redundant mitochondria (Fig. 1), not disposed of by autophagy, can represent a threat to cellular homeostasis and cell survival. In the yeast, Saccharomyces cerevisiae, nutrient deprivation (e.g., nitrogen starvation) or damage can promote selective turnover of mitochondria by autophagy in a process termed mitophagy 5-9.
We describe a simple fluorescence microscopy approach to assess autophagy. For clarity we restrict our description here to show how the approach can be used to monitor mitophagy in yeast cells. The assay makes use of a fluorescent reporter, Rosella, which is a dual-emission biosensor comprising a relatively pH-stable red fluorescent protein linked to a pH-sensitive green fluorescent protein. The operation of this reporter relies on differences in pH between the vacuole (pH ~ 5.0-5.5) and mitochondria (pH ~ 8.2) in living cells. Under growing conditions, wild type cells exhibit both red and green fluorescence distributed in a manner characteristic of the mitochondria. Fluorescence emission is not associated with the vacuole. When subjected to nitrogen starvation, a condition which induces mitophagy, in addition to red and green fluorescence labeling the mitochondria, cells exhibit the accumulation of red, but not green fluorescence, in the acidic vacuolar lumen representing the delivery of mitochondria to the vacuole. Scoring cells with red, but not green fluorescent vacuoles can be used as a measure of mitophagic activity in cells5,10-12.

A Fluorescence Microscopy Assay for Monitoring Mitophagy in the Yeast Saccharomyces cerevisiae

A robust approach to monitor the delivery of organelles to the acidic lumen of the yeast vacuole for degradation and recycling is described. The method relies on the specific labeling of target organelles with a genetically encoded dual-emission fluorescence pH-biosensor, and visualization of individual cells using fluorescence microscopy.

Maya İzleme Mitophagy Floresan Mikroskopi Testi Saccharomyces cerevisiae

Autophagy is important for turnover of cellular components under a range of different conditions. It serves an essential homeostatic function as well as a ...

Biyoloji

Quantification of Efferocytosis by Single-cell Fluorescence Microscopy

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling and cellular processes that control efferocytosis. This quantification can be difficult to perform as apoptotic cells are often efferocytosed piecemeal, thus necessitating methods which can accurately delineate between the efferocytosed portion of an apoptotic target versus residual unengulfed cellular fragments. The approach outlined herein utilizes dual-labeling approaches to accurately quantify the dynamics of efferocytosis and efferocytic capacity of efferocytes such as macrophages. The cytosol of the apoptotic cell is labeled with a cell-tracking dye to enable monitoring of all apoptotic cell-derived materials, while surface biotinylation of the apoptotic cell allows for differentiation between internalized and non-internalized apoptotic cell fractions. The efferocytic capacity of efferocytes is determined by taking fluorescent images of live or fixed cells and quantifying the amount of bound versus internalized targets, as differentiated by streptavidin staining. This approach offers several advantages over methods such as flow cytometry, namely the accurate delineation of non-efferocytosed versus efferocytosed apoptotic cell fractions, the ability to measure efferocytic dynamics by live-cell microscopy, and the capacity to perform studies of cellular signaling in cells expressing fluorescently-labeled transgenes. Combined, the methods outlined in this protocol serve as the basis for a flexible experimental approach that can be used to accurately quantify efferocytic activity and interrogate cellular signaling pathways active during efferocytosis.

Efferocytosis, the phagocytic removal of apoptotic cells, is required to maintain homeostasis and is facilitated by receptors and signaling pathways that allow for the recognition, engulfment, and internalization of apoptotic cells. Herein, we present a fluorescence microscopy protocol for the quantification of efferocytosis and the activity of efferocytic signaling pathways.

Efferocytosis tek hücreli floresans mikroskobu tarafından miktar

Studying the regulation of efferocytosis requires methods that are able to accurately quantify the uptake of apoptotic cells and to probe the signaling ...

İmmünoloji ve Enfeksiyon

A Faster, High Resolution, mtPA-GFP-based Mitochondrial Fusion Assay Acquiring Kinetic Data of Multiple Cells in Parallel Using Confocal Microscopy

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in living cells by photo-conversion of matrix targeted photoactivatable GFP (mtPAGFP) in a subset of mitochondria. The rate at which the photoconverted molecules equilibrate across the entire mitochondrial population is used as a measure of fusion activity. Thus far measurements were performed using a single cell time lapse approach, quantifying the equilibration in one cell over an hour. Here, we scale up and automate a previously published live cell method based on using mtPAGFP and a low concentration of TMRE (15 nm). This method involves photoactivating a small portion of the mitochondrial network, collecting highly resolved stacks of confocal sections every 15 min for 1 hour, and quantifying the change in signal intensity. Depending on several factors such as ease of finding PAGFP expressing cells, and the signal of the photoactivated regions, it is possible to collect around 10 cells within the 15 min intervals. This provides a significant improvement in the time efficiency of this assay while maintaining the highly resolved subcellular quantification as well as the kinetic parameters necessary to capture the detail of mitochondrial behavior in its native cytoarchitectural environment.
Mitochondrial dynamics play a role in many cellular processes including respiration, calcium regulation, and apoptosis1,2,3,13. The structure of the mitochondrial network affects the function of mitochondria, and the way they interact with the rest of the cell. Undergoing constant division and fusion, mitochondrial networks attain various shapes ranging from highly fused networks, to being more fragmented. Interestingly, Alzheimer's disease, Parkinson's disease, Charcot Marie Tooth 2A, and dominant optic atrophy have been correlated with altered mitochondrial morphology, namely fragmented networks4,10,13. Often times, upon fragmentation, mitochondria become depolarized, and upon accumulation this leads to impaired cell function18. Mitochondrial fission has been shown to signal a cell to progress toward apoptosis. It can also provide a mechanism by which to separate depolarized and inactive mitochondria to keep the bulk of the network robust14. Fusion of mitochondria, on the other hand, leads to sharing of matrix proteins, solutes, mtDNA and the electrochemical gradient, and also seems to prevent progression to apoptosis9. How fission and fusion of mitochondria affects cell homeostasis and ultimately the functioning of the organism needs further understanding, and therefore the continuous development and optimization of how to gather information on these phenomena is necessary.
Existing mitochondrial fusion assays have revealed various insights into mitochondrial physiology, each having its own advantages. The hybrid PEG fusion assay7, mixes two populations of differently labeled cells (mtRFP and mtYFP), and analyzes the amount of mixing and colocalization of fluorophores in fused, multinucleated, cells. Although this method has yielded valuable information, not all cell types can fuse, and the conditions under which fusion is stimulated involves the use of toxic drugs that likely affect the normal fusion process. More recently, a cell free technique has been devised, using isolated mitochondria to observe fusion events based on a luciferase assay1,5. Two human cell lines are targeted with either the amino or a carboxy terminal part of Renilla luciferase along with a leucine zipper to ensure dimerization upon mixing. Mitochondria are isolated from each cell line, and fused. The fusion reaction can occur without the cytosol under physiological conditions in the presence of energy, appropriate temperature and inner mitochondrial membrane potential. Interestingly, the cytosol was found to modulate the extent of fusion, demonstrating that cell signaling regulates the fusion process 4,5. This assay will be very useful for high throughput screening to identify components of the fusion machinery and also pharmacological compounds that may affect mitochondrial dynamics. However, more detailed whole cell mitochondrial assays will be needed to complement this in vitro assay to observe these events within a cellular environment.
A technique for monitoring whole-cell mitochondrial dynamics has been in use for some time and is based on a mitochondrially-targeted photoactivatable GFP (mtPAGFP)6,11. Upon expression of the mtPAGFP, a small portion of the mitochondrial network is photoactivated (10-20%), and the spread of the signal to the rest of the mitochondrial network is recorded every 15 minutes for 1 hour using time lapse confocal imaging. Each fusion event leads to a dilution of signal intensity, enabling quantification of the fusion rate. Although fusion and fission are continuously occurring in cells, this technique only monitors fusion as fission does not lead to a dilution of the PAGFP signal6. Co-labeling with low levels of TMRE (7-15 nM in INS1 cells) allows quantification of the membrane potential of mitochondria. When mitochondria are hyperpolarized they uptake more TMRE, and when they depolarize they lose the TMRE dye. Mitochondria that depolarize no longer have a sufficient membrane potential and tend not to fuse as efficiently if at all. Therefore, active fusing mitochondria can be tracked with these low levels of TMRE9,15. Accumulation of depolarized mitochondria that lack a TMRE signal may be a sign of phototoxicity or cell death. Higher concentrations of TMRE render mitochondria very sensitive to laser light, and therefore great care must be taken to avoid overlabeling with TMRE. If the effect of depolarization of mitochondria is the topic of interest, a technique using slightly higher levels of TMRE and more intense laser light can be used to depolarize mitochondria in a controlled fashion (Mitra and Lippincott-Schwartz, 2010). To ensure that toxicity due to TMRE is not an issue, we suggest exposing loaded cells (3-15 nM TMRE) to the imaging parameters that will be used in the assay (perhaps 7 stacks of 6 optical sections in a row), and assessing cell health after 2 hours. If the mitochondria appear too fragmented and cells are dying, other mitochondrial markers, such as dsRED or Mitotracker red could be used instead of TMRE.
The mtPAGFP method has revealed details about mitochondrial network behavior that could not be visualized using other methods. For example, we now know that mitochondrial fusion can be full or transient, where matrix content can mix without changing the overall network morphology. Additionally, we know that the probability of fusion is independent of contact duration and organelle dimension, is influenced by organelle motility, membrane potential and history of previous fusion activity8,15,16,17.
In this manuscript, we describe a methodology for scaling up the previously published protocol using mtPAGFP and 15nM TMRE8 in order to examine multiple cells at a time and improve the time efficiency of data collection without sacrificing the subcellular resolution. This has been made possible by the use of an automated microscope stage, and programmable image acquisition software. Zen software from Zeiss allows the user to mark and track several designated cells expressing mtPAGFP. Each of these cells can be photoactivated in a particular region of interest, and stacks of confocal slices can be monitored for mtPAGFP signal as well as TMRE at specified intervals. Other confocal systems could be used to perform this protocol provided there is an automated stage that is programmable, an incubator with CO2, and a means by which to photoactivate the PAGFP; either a multiphoton laser, or a 405 nm diode laser.

Mitochondrial fusion was measured by tracking the equilibration of photoconverted matrix-targeted GFP across the mitochondrial network over time. Thus far, only one cell could be subjected to an hour long kinetic analysis at a time. We present a method that simultaneously measures multiple cells, thereby speeding up the data collection process.

Paralel Konfokal Mikroskop kullanarak birden fazla Hücre Kinetik Veri Kazanılması A Daha hızlı, Yüksek Çözünürlük, MTPA-GFP tabanlı Mitokondrial Füzyon Deneyi

Mitochondrial fusion plays an essential role in mitochondrial calcium homeostasis, bioenergetics, autophagy and quality control. Fusion is quantified in ...

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a wide variety of disorders including metabolic diseases, premature aging syndromes, and neurodegenerative diseases such as Alzheimer's disease (AD) and Parkinson's disease (PD). Maintenance of mitochondrial health depends on mitochondrial biogenesis and the efficient clearance of dysfunctional mitochondria through mitophagy. Experimental methods to accurately detect autophagy/mitophagy, especially in animal models, have been challenging to develop. Recent progress towards the understanding of the molecular mechanisms of mitophagy has enabled the development of novel mitophagy detection techniques. Here, we introduce several versatile techniques to monitor mitophagy in human cells, Caenorhabditis elegans (e.g., Rosella and DCT-1/ LGG-1 strains), and mice (mt-Keima). A combination of these mitophagy detection techniques, including cross-species evaluation, will improve the accuracy of mitophagy measurements and lead to a better understanding of the role of mitophagy in health and disease.

In Vitro and In Vivo Detection of Mitophagy in Human Cells, C. Elegans, and Mice

Mitophagy, the process of clearing damaged mitochondria, is necessary for mitochondrial homeostasis and health maintenance. This article presents some of the latest mitophagy detection methods in human cells, Caenorhabditis elegans, and mice.

Vitro ve In Vivo insan hücreleri, C. Elegansve fareler Mitophagy tespiti

Mitochondria are the powerhouses of cells and produce cellular energy in the form of ATP. Mitochondrial dysfunction contributes to biological aging and a ...

Tıp

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

Ateş Drosophila ImageJ kullanarak Fluorescently Tagged proteinlerin tarafsız analizi ile yılında niceliksel hücre biyolojisi

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...

Nörobilim

Visualization of Endogenous Mitophagy Complexes In Situ in Human Pancreatic Beta Cells Utilizing Proximity Ligation Assay

Mitophagy is an essential mitochondrial quality control pathway, which is crucial for pancreatic islet beta cell bioenergetics to fuel glucose-stimulated insulin release. Assessment of mitophagy is challenging and often requires genetic reporters or multiple complementary techniques not easily utilized in tissue samples, such as primary human pancreatic islets. Here we demonstrate a robust approach to visualize and quantify formation of key endogenous mitophagy complexes in primary human pancreatic islets. Utilizing the sensitive proximity ligation assay technique to detect interaction of the mitophagy regulators NRDP1 and USP8, we are able to specifically quantify formation of essential mitophagy complexes in situ. By coupling this approach to counterstaining for the transcription factor PDX1, we can quantify mitophagy complexes, and the factors that can impair mitophagy, specifically within beta cells. The methodology we describe overcomes the need for large quantities of cellular extracts required for other protein-protein interaction studies, such as immunoprecipitation (IP) or mass spectrometry, and is ideal for precious human islet&#160;samples generally not available in sufficient quantities for these approaches. Further, this methodology obviates the need for flow sorting techniques to purify beta cells from a heterogeneous islet population for downstream protein applications. Thus, we describe a valuable protocol for visualization of mitophagy highly compatible for use in heterogeneous and limited cell populations.

This protocol outlines a method for quantitative analysis of mitophagy protein complex formation specifically in beta cells from primary human islet samples. This technique thus allows analysis of mitophagy from limited biological material, which are crucial in precious human pancreatic beta cell samples.

Yakınlık ligasyon assay kullanarak ınsan pankreatik Beta hücrelerinde situ Içinde endojen Mitophagy kompleksleri görselleştirme

Mitophagy is an essential mitochondrial quality control pathway, which is crucial for pancreatic islet beta cell bioenergetics to fuel glucose-stimulated ...

Sensitive Measurement of Mitophagy by Flow Cytometry Using the pH-dependent Fluorescent Reporter mt-Keima

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between defective mitophagy and various human diseases, including neurodegenerative diseases, cancer, and metabolic diseases. In addition, recent studies have shown that mitophagy is involved in normal cellular processes, such as differentiation and development. However, the precise role of and molecular mechanisms underlying mitophagy require further study. Therefore, it is critical to develop a robust and convenient method for measuring changes in mitophagy activity. Here, we describe a detailed protocol for quantitatively assessing mitophagy activity through flow cytometry using the mitochondria-targeted fluorescent protein Keima (mt-Keima). This flow cytometry assay can analyze mitophagy activity more rapidly and sensitively than conventional microscopy- or immunoblotting-based methods. This protocol can be applied to analyze mitophagy activity in various cell types.

Mitophagy, the selective degradation of mitochondria, has been implicated in mitochondrial homeostasis and is deregulated in various human diseases. However, convenient experimental methods for measuring mitophagy activity are very limited. Here, we provide a sensitive assay for measuring mitophagy activity using flow cytometry.

Hassas ölçüm Mitophagy floresan muhabir akış sitometresi kullanarak pH bağımlı mt-Keima tarafından

Mitophagy is a process of selective removal of damaged or unnecessary mitochondria using autophagy machinery. Close links have been found between ...

Protection of H9c2 Myocardial Cells from Oxidative Stress by Crocetin via PINK1/Parkin Pathway-Mediated Mitophagy

This study aimed to explore the oxidative stress-protective effect of crocetin on H2O2-mediated H9c2 myocardial cells through in vitro experiments, and further explore whether its mechanism is related to the impact of mitophagy. This study also aimed to demonstrate the therapeutic effect of safflower acid on oxidative stress in cardiomyocytes and explore whether its mechanism is related to the effect of mitophagy. Here, an H2O2-based oxidative stress model was constructed and assessed the degree of oxidative stress injury of cardiomyocytes by detecting the levels of lactate dehydrogenase (LDH), creatine kinase (CK), malondialdehyde (MDA), superoxide dismutase (SOD), catalase (CAT), and glutathione peroxidase (GSH Px). Reactive oxygen species (ROS)-detecting fluorescent dye DCFH-DA, JC-1 dye, and TUNEL dye were employed to assess mitochondrial damage and apoptosis. Autophagic flux was measured by transfecting Ad-mCherry-GFP-LC3B adenovirus. Mitophagy-related proteins were then detected via western blotting and immunofluorescence. However, crocetin (0.1-10 &#181;M) could significantly improve cell viability and reduce apoptosis and oxidative stress damage caused by H2O2. In cells with excessive autophagic activation, crocetin could also reduce autophagy flow and the expression of mitophagy-related proteins PINK1 and Parkin, and reverse the transfer of Parkin to mitochondria. Crocetin could reduce H2O2-mediated oxidative stress damage and the apoptosis of H9c2 cells, and its mechanism was closely related to mitophagy.

Protection of H9c2 Myocardial Cells from Oxidative Stress by Crocetin via PINK1/Parkin Pathway-Mediated Mitophagy

Based on in vitro experiments, this study revealed the mechanism of crocetin in repairing oxidative stress damage of cardiomyocytes by influencing mitophagy, in which the PINK1/Parkin signaling pathway plays an important role.

H9c2 Miyokard Hücrelerinin Crocetin ile Oksidatif Stresten PINK1 / Parkin Yolu Aracılı Mitopaji ile Korunması

This study aimed to explore the oxidative stress-protective effect of crocetin on H2O2-mediated H9c2 myocardial cells through in vitro experiments, and ...

Visualizing Mitophagy with Fluorescent Dyes for Mitochondria and Lysosome

Mitochondria, being the powerhouses of the cell, play important roles in bioenergetics, free radical generation, calcium homeostasis, and apoptosis. Mitophagy is the primary mechanism of mitochondrial quality control and is generally studied using microscopic observation, however in vivo mitophagy assays are difficult to perform. Evaluating mitophagy by imaging live organelles is an alternative and necessary method for mitochondrial research. This protocol describes the procedures for using the cell-permeant green-fluorescent mitochondria dye MitoTracker Green and the red-fluorescent lysosome dye LysoTracker Red in live cells, including the loading of the dyes, visualization of the mitochondria and the lysosome, and expected outcomes. Detailed steps for the evaluation of mitophagy in live cells, as well as technical notes about microscope software settings, are also provided. This method can help researchers observe mitophagy using live-cell fluorescent microscopy. In addition, it can be used to quantify mitochondria and lysosomes and assess mitochondrial morphology.

Mitophagy is the primary mechanism of mitochondrial quality control. However, the evaluation of mitophagy in vivo is hindered by the lack of reliable quantitative assays. Presented here is a protocol for the observation of mitophagy in living cells using a cell-permeant green-fluorescent mitochondria dye and a red-fluorescent lysosome dye.

Mitokondri ve Lizozom için Floresan Boyalarla Mitopajinin Görselleştirilmesi

Mitochondria, being the powerhouses of the cell, play important roles in bioenergetics, free radical generation, calcium homeostasis, and apoptosis. ...

Mitofajinin Ölçülmesi C. elegans ve Karaciğer Kanseri Hücre Hattı

Detection of Mitophagy in Caenorhabditis elegans and Mammalian Cells Using Organelle-Specific Dyes

Mitochondria are essential for various biological functions, including energy production, lipid metabolism, calcium homeostasis, heme biosynthesis, regulated cell death, and the generation of reactive oxygen species (ROS). ROS are vital for key biological processes. However, when uncontrolled, they can lead to oxidative injury, including mitochondrial damage. Damaged mitochondria release more ROS, thereby intensifying cellular injury and the disease state. A homeostatic process named mitochondrial autophagy (mitophagy) selectively removes damaged mitochondria, which are then replaced by new ones. There are multiple mitophagy pathways, with the common endpoint being the breakdown of the damaged mitochondria in lysosomes. 
Several methodologies, including genetic sensors, antibody immunofluorescence, and electron microscopy, use this endpoint to quantify mitophagy. Each method for examining mitophagy has its advantages, such as specific tissue/cell targeting (with genetic sensors) and great detail (with electron microscopy). However, these methods often require expensive resources, trained personnel, and a lengthy preparation time before the actual experiment, such as for creating transgenic animals. Here, we present a cost-effective alternative for measuring mitophagy using commercially available fluorescent dyes targeting mitochondria and lysosomes. This method effectively measures mitophagy in the nematode Caenorhabditis elegans and human liver cells, which indicates its potential efficiency in other model systems.

Yazar Gündemi: Organele Özgü Boyalar Kullanılarak Caenorhabditis elegans ve Memeli Hücrelerinde Mitofajinin Tespiti  

Exploring mitophagy through electron microscopy, genetic sensors, and immunofluorescence requires costly equipment, skilled personnel, and a significant time investment. Here, we demonstrate the efficacy of a commercial fluorescence dye kit in quantifying the mitophagy process in both Caenorhabditis elegans and a liver cancer cell line.

Caenorhabditis elegans ve Memeli Hücrelerinde Mitopajinin Organele Özgü Boyalar Kullanılarak Saptanması

Mitochondria are essential for various biological functions, including energy production, lipid metabolism, calcium homeostasis, heme biosynthesis, ...