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How come Draq5 stains cytoplasm and membranes under confocal

How come Draq5 stains cytoplasm and membranes under confocal


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Just an imaging question on the back of my head. For nuclear counter stain, Draq5 has been an excellent choice for many people over many years. And I too love it.

The working principle of Draq5 is the nuclear dye Draq5 binds to doubles strand DNA, especially d-A and d-T. That is to say it is very specific to stain nucleus DNA, minimal binding to mitochondria DNA, and no binding to RNA. It is also a very good live cell permeant dye.

In practice, we use it to stain cell nucleus and then perform image analysis, the primary objective in image analysis is identification of the cell object, usually through nucleus staining. No problem, Draq5 does this perfectly, I can see my image crispy clear.

Next, it comes to the estimation of the entire cell, that is to find cell membrane boundaries. A huge advantage of Draq 5 is it also stain cytoplasm and cell membrane weakly, allowing the accurate detection of cell boundaries. Comparing this to Hoechst dye, which also stain nucleus and weakly cytoplasm only for estimation of cell boundaries. This is unlikely a Draq5 non-specific binding, since data from resistant cancer cells that have membrane drug pumps still retain Draq5 very well.

I'm not a chemist so that one thing in the back of my head is: Why does Draq5 bind cytoplasm and outlines cell membranes that well? Given it is highly specific to bind A/T in double strand DNA grooves. Presumably the dye is binding to its cytoplasm target and membrane target via different mechanisms. Can someone explain that?


Discrimination of DNA and RNA in cells by a vital fluorescent probe: Lifetime imaging of SYTO13 in healthy and apoptotic cells

Of the few vital DNA and RNA probes, the SYTO dyes are the most specific for nucleic acids. However, they show no spectral contrast upon DNA or RNA binding. We show that fluorescence lifetime imaging using two-photon excitation of SYTO13 allows differential and simultaneous imaging of DNA and RNA in living cells, as well as sequential and repetitive assessment of staining patterns.

Methods

Two-photon imaging of SYTO13 is combined with lifetime contrast, using time-gated detection. We focus on distinguishing DNA and RNA in healthy and apoptotic Chinese hamster ovary cells.

Results

In healthy cells, SYTO13 has a fluorescence lifetime of 3.4 ± 0.2 ns when associated with nuclear DNA. Bound to RNA, its lifetime is 4.1 ± 0.1 ns. After induction of apoptosis, clusters of SYTO13 with fluorescence lifetime of 3.4 ± 0.2 ns become apparent in the cytoplasm. They are identified as mitochondrial DNA on the basis of colocalization experiments with the DNA-specific dye, DRAQ5, and the mitochondrial-specific dye, CMXRos. Upon progression of apoptosis, the lifetime of SYTO13 attached to DNA shortens significantly, which is indicative of changes in the molecular environment of the dye.

Conclusions

We have characterized SYTO13 as a vital lifetime probe, allowing repetitive and differential imaging of DNA and RNA. Cytometry 47:226–235, 2002. © 2002 Wiley-Liss, Inc.

Simultaneous and differential imaging of dynamic properties of DNA and RNA in various compartments of vital cells is of interest for a number of reasons. For example, visualization of the compartmentalization of transcriptional activity in the nucleus of living cells is still difficult with the nucleic acid probes available ( 1 ). These dyes often exhibit similar affinities for DNA and RNA. In addition, the aggregation, cleavage, and segregation of DNA in the apoptotic nucleus ( 2 ) cannot be studied properly in living cells in combination with changes in RNA structure. This is because no vital imaging probes are available that distinguish nuclear DNA from nucleolar RNA or from mitochondrial DNA. Therefore, segregation of nuclear DNA and RNA to the cytoplasm is difficult to monitor. Studying the dynamics of nucleic acid behavior in living cells will benefit greatly from probes or methods that allow the separate, but simultaneous, detection of DNA and RNA ( 3-5 ).

Spectroscopic or lifetime contrast may be used to distinguish the binding sites of a fluorescent probe. In spectroscopic imaging ( 6 , 7 ) differences in fluorescence intensity at two excitation or various emission wavelengths are detected. In lifetime imaging ( 8-14 ), the typical time constant of the fluorescence decay processes is used as the contrast mechanism. Both principles can be applied for the simultaneous imaging of multiple probes or for a single probe that exhibits different spectral or lifetime properties, depending on its binding partner.

Because only few vital imaging probes are available that bind exclusively to either DNA or RNA, the use of a single probe is preferred. Probes of the SYTO family ( 15 , 16 ) are cell-permeable nucleic acid stains with varying characteristics, such as cell permeability, fluorescence enhancement upon binding, excitation and emission spectra, and DNA/RNA binding affinity. However, they exhibit no spectral contrast in binding to DNA or RNA. Therefore, the question arises: Do the SYTO probes exhibit lifetime contrast on binding sites?

The fluorescence lifetime of a probe is independent of characteristics that do modulate the detected fluorescence intensity (e.g., laser intensity fluctuations, photobleaching, probe leakage, out-of-focus movements). On the other hand, it can be sensitive to chemical parameters in its environment, like pH ( 14 , 17-19 ) and ion or oxygen concentrations ( 12 , 20-27 ). It may yield quantitative information about the local chemical environment and state of the fluorescent molecules in the cell ( 13 , 14 , 28-30 ) and therefore offer a tool to discriminate between DNA and RNA-bound SYTO dyes.

We applied a combination ( 31 ) of fluorescence lifetime imaging and two-photon excitation to study the effect of binding of DNA and RNA on the properties of one of the SYTO probes, i.e., SYTO13. Two-photon excitation microscopy ( 32 ) is now used widely in life sciences ( 33-36 ). It not only has an intrinsically high axial resolution ( 37 ) without the need of a confocal pinhole in the detection path, but its use can also significantly reduce cell damage, both due to the use of longer wavelength excitation light (avoiding the use of damaging UV or blue excitation light) and the reduction of out-of-focus irradiation ( 38 , 39 ). This makes two-photon fluorescence microscopy very suitable for the repetitive imaging of cells ( 35 , 39 ), without seriously affecting their vitality. Because two-photon microscopy requires the use of femtosecond pulsed lasers ( 32 ), the combination of two-photon and lifetime contrast imaging with the application of the time-gated detection technique ( 14 ) is straightforward.

SYTO13 ( 15 ) becomes fluorescent when bound to DNA or RNA. The quantum yields are equal when bound to RNA and DNA, its excitation spectrum has a maximum at 488 nm, and the emission spectrum peaks at 510 nm. Also, the binding affinity characteristics for DNA and RNA are comparable. Finally, SYTO13 labels DNA, both in the nucleus and in the mitochondria, and RNA in the cytoplasm and nucleoli. Our study was carried out in healthy Chinese hamster ovary (CHO) cells. Based on earlier observations that some DNA probes show changes in fluorescence lifetime properties upon apoptosis ( 40-42 ), we extended the study to cells that were induced to enter programmed cell death. Where necessary, observations were verified by dual-labeling experiments with probes of known labeling properties, i.e., DRAQ5 (a DNA probe 43) and CMXRos (a mitochondrial probe 15,44). Colocalization was present when energy transfer, as visualized by changes in the lifetime of SYTO13 fluorescence, occurred between both dyes or when structures coincided. We show that two-photon fluorescence lifetime imaging of SYTO13 has potential for vital and differential imaging of DNA and RNA dynamics without damaging effects to the cells.


MATERIALS AND METHODS

Cell Cultures

CHO cells were grown on coverslips in HAM's F12 medium (ICN Biomedicals, Costa Mesa, CA) supplemented with 10% fetal calf serum (FCS Gibco Life Technologies, Paisley, UK) and 2 mM L-glutamine. Cell cultures were maintained in an incubator at 5% CO2 and 37°C. Prior to apoptosis, cells were maintained overnight in HAM's F12 medium supplemented with 1% FCS and 2 mM L-glutamine. Apoptosis was induced by the addition of staurosporin (1 μM final concentration, Sigma, Zwijndrecht, The Netherlands) 2 h before the imaging experiments started.

Labeling Procedures and Sample Preparation

To determine the lifetime characteristics of SYTO13 interacting with DNA or RNA, independent of its cellular environment, isolated DNA and tRNA were labeled with SYTO13 by the addition of the probe to DNA or tRNA in phosphate-buffered saline (PBS) to a final concentration of 2 μM. Yeast-tRNA (10 mg/ml) was obtained from Roche Molecular Biochemicals (Basel, Switzerland), whereas salmon testis DNA (10 mg/ml) was obtained from Sigma. SYTO13 (in dimethylsulfoxide [DMSO], 5mM) and CMXRos (in methanol, 5 mM) were obtained from Molecular Probes (Leiden, The Netherlands). DRAQ5 (5 mM in HCl) was a gift from Prof. dr. Paul J. Smith (Department of Pathology, University of Wales, College of Medicine, Cardiff, UK).

Healthy and apoptotic cells were labeled by the addition of SYTO13-DMSO (final concentration 1 μM) to the cells on coverslips in medium. After 30 min of incubation, SYTO13 was washed away with probe-free medium and images were taken. Labeling of CHO cells with SYTO13 and repetitive imaging of the cells did not induce any morphological changes.

Using the standard labeling procedure ( 43 ) for the DNA probe, DRAQ5, cells die rapidly, presumably by necrosis. Therefore, an alternative staining procedure was developed. Healthy and apoptotic cells were stained with DRAQ5 by the addition of the probe to cells on coverslips in medium to a final concentration of 1 μM. After only 3 min, the staining medium was removed and replaced by fresh, probe-free medium. Applying this procedure, cells exhibited no morphological changes within the time span of the imaging experiments.

Dual labeling of cells with both SYTO13 and DRAQ5 was carried out by first labeling with SYTO13, followed by the addition of DRAQ5 according to the procedure described above. After only 3 min, the staining medium containing both SYTO13 and DRAQ5 was removed and replaced by probe-free medium.

The mitochondria of apoptotic cells were labeled for 1 h by addition of CMXRos ( 15 ) to cells on coverslips in medium to a final concentration of 0.5 mM, prior to the addition of staurosporin. In dual-labeling experiments of CMXRos and SYTO13, the latter was added to cells already labeled with CMXRos prior to observation under the microscope, using the SYTO13 labeling procedure described above. Coverslips with either cells or droplets of isolated DNA/RNA solutions were mounted in a home-built sample holder and viewed under the microscope.

Two-Photon Lifetime Microscopy

Experiments were carried out on a home-built two-photon excitation inverted microscope as described in detail elsewhere ( 31 ). In short, the system is equipped with a mode-locked titanium:sapphire (Ti:Sa) laser (Tsunami, Spectra-Physics, Mountain View, CA) that produces 80-fs pulses at a repetition rate of 82 MHz. After passing a beam expander and neutral density filters, the excitation light is directed to a dichroic mirror that is highly reflective in the near infrared and transparent below 700 nm. Thus, the excitation light is reflected toward and focused by a 60× plan-apochromatic water-immersion objective (NA1.2, Nikon/Uvikon, Bunnik, The Netherlands) and scanned over the sample. The fluorescence light is collected by the same objective. The optical resolution was 0.27 μm laterally and 0.72 μm axially at an excitation wavelength of 800 nm ( 45 ). All fluorescence light below 700 nm is transmitted through the dichroic mirror described above and collected by a photomultiplier (Hamamatsu R1894, Toyooka Village, Japan) in photon counting mode. Additional blocking of excitation light is achieved by means of a series of 750-nm interference short pass filters in the imaging path behind the mirror. If desired, a more narrow emission wavelength range can be selected by placing additional interference (IF) or long pass (LP) filters (Optosigma, Santa Anna, CA) in the emission pathway.

All fluorescence (lifetime) imaging experiments were carried out at an excitation wavelength of 800 nm. The fluorescence emission after pulsed excitation can be described by an exponential decay with a typical lifetime. Fluorescence lifetimes of commonly used dyes are typically on the order of a few nanoseconds. The method used here to determine the fluorescence lifetime is based on time-gated detection of the fluorescence. After each excitation pulse, the fluorescence decay is detected in four time-windows (gates) that are enabled sequentially, making the acquired lifetime intrinsically independent of laser intensity fluctuations. Details of the lifetime module are given elsewhere ( 14 ). The signal from many excitation pulses is integrated for each pixel. The integrated intensity in the four time gates is fitted to a monoexponential decay using a nonlinear least square Levenberg-Marquardt fit algorithm ( 46 ). The accumulated intensity of the four gates yields the fluorescence intensity for each pixel of the specimen. The fitted lifetime for every pixel results in the corresponding lifetime image. Error margins (SEM) are determined from the fit results within an area (10 × 10 pixels) of similar lifetimes. Good quality fluorescence lifetime images of 256 × 256 pixels are recorded in about 30 s. All images shown are single-section images of a cell, thus utilizing the high axial resolution of two-photon excitation. In general, areas of short lifetime and high intensity are present next to areas of short lifetime and low intensity. A similar observation is made for long lifetimes. This indicates that the observed differences in lifetime are generally not correlated with intensity variations due to bleaching or differences in local probe concentrations. Data were processed using home-written programs under the IDL software package (Creaso B.V., Apeldoorn, The Netherlands).

Fluorescence Spectra of SYTO13, DRAQ5, and CMXRos

Two-photon excitation spectra may differ (in shape and peak wavelengths) from a doubled-wavelength version of the one-photon excitation spectra because the selection rules are different ( 36 ). Although the one-photon excitation spectra of the probes used here differ significantly from each other ( 15 , 43 ), all appear to be excitable with two-photon excitation at 800 nm. For CMXRos and DRAQ5, the obtained fluorescence intensity is relatively low.

Fluorophores show the same emission spectrum with either mode of excitation ( 36 ). Figure 1 shows the emission spectrum of SYTO13-DNA ( 15 ), CMXRos-methanol ( 15 ), and DRAQ5-PBS in solutions. Spectral measurements on DRAQ5 diluted in PBS (1 μM) were carried out on an SPF-8100 series 2 fluorometer (SLM-Aminco, Beun de Ronde, Culemborg, The Netherlands). For determination of the emission spectrum, the probe was excited at 568 nm. For the excitation spectrum, the emitted fluorescence was detected at 670 nm. In cells, the emission spectra of SYTO13, CMXRos, and DRAQ5 were measured using a spectral microscope ( 7 ) with excitation at 488 nm for SYTO13 and CMXRos and 568 nm for DRAQ5 (spectra not shown). We found that the emission spectra of SYTO13 and CMXRos are unaltered in cells, whereas the emission spectrum of DRAQ5 shows a significant red shift of 20 nm, in agreement with earlier observations ( 43 ). Apoptosis did not have an influence on the shape or peak positions of the emission spectra for any of the probes.

Emission spectra of the probes SYTO13-DNA (⧫, excitation at 488 nm), CMXRos-methanol (▪, excitation at 488 nm), and DRAQ5-PBS (•, excitation at 568 nm). Also indicated (-⧫-) is the position of the 800-nm excitation pulse in the two-photon experiments.

Autofluorescence of CHO Cells

We determined the intensity, wavelength range, and the lifetimes of the autofluorescence of healthy and apoptotic cells and compared them with the fluorescence characteristics of the various probes. Upon two-photon excitation with 800 nm, both healthy and apoptotic CHO cells exhibit weak autofluorescence within the wavelength range of 500–600 nm. No autofluorescence was detected above 600 nm. No differences in autofluorescence distribution and lifetimes between healthy and apoptotic cells were seen. Autofluorescence is distributed homogeneously over the cytoplasm, with occasionally scattered spots of higher intensity. The lifetime of the autofluorescence varies: 2.5 ± 0.4 ns (diffuse) and 4.0 ± 0.5 ns (spots). In an earlier study ( 34 ), the value of 2.2 ns was attributed to the fluorescence of NADH binding to proteins (e.g., in the mitochondria).

Use of Emission Filters in Single and Double-Probe Experiments

On the basis of the overlap of the spectral region of cellular autofluorescence and the emission spectra of SYTO13 (for the latter, see Fig. 1), the use of an emission filter (IF530 ± 60) nm for the suppression of autofluorescence relative to SYTO13 fluorescence is not possible. However, because the autofluorescence is very weak, a specific set-up configuration could be found (e.g., excitation light intensity, dwell time) in which autofluorescence did not exceed the detection limit and thus did not contribute to the fluorescence signal of SYTO13. In contrast, for the weakly fluorescent probes DRAQ5 and CMXRos, the filters LP650 and IF600 ± 30 nm had to be used, respectively, to suppress the contribution of autofluorescence.

In the colocalization experiments, cells were labeled simultaneously with two probes (SYTO13/DRAQ5 and SYTO13/CMXRos). In these pairs, the probes have different emission characteristics (Fig. 1) and, therefore, the emission filters given above can be used to separate their fluorescence. Sequential observation through either of the filters now allows colocalization of the probes.


How come Draq5 stains cytoplasm and membranes under confocal - Biology

Recent work has underscored the importance of membrane trafficking events during cytokinesis. For example, targeted membrane secretion occurs at the cleavage furrow in animal cells, and proteins that regulate endocytosis also influence the process of cytokinesis. Nonetheless, the prevailing dogma is that endosomal membrane trafficking ceases during mitosis and resumes after cell division is complete. In this study, we have characterized endocytic membrane trafficking events that occur during mammalian cell cytokinesis. We have found that, although endocytosis ceases during the early stages of mitosis, it resumes during late mitosis in a temporally and spatially regulated pattern as cells progress from anaphase to cytokinesis. Using fixed and live cell imaging, we have found that, during cleavage furrow ingression, vesicles are internalized from the polar region and subsequently trafficked to the midbody area during later stages of cytokinesis. In addition, we have demonstrated that cytokinesis is inhibited when clathrin-mediated endocytosis is blocked using a series of dominant negative mutants. In contrast to previous thought, we conclude that endocytosis resumes during the later stages of mitosis, before cytokinesis is completed. Furthermore, based on our findings, we propose that the proper regulation of endosomal membrane traffic is necessary for the successful completion of cytokinesis.

This work was supported in part by grants from the United States Department of Defense and the American Cancer Society (RSG 03-023-01-CSM) (to C. D-S.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. 1-7, of which supplemental Figs. 3-6 consist of both still images and videos.

Recipient of a Special Fellow Career Development Award from The Leukemia and Lymphoma Society.


Once dry the microorganisms can be seen under a microscope. This procedure is very different from that of a differential stain. A common differential stain a.

What is meant by quantitative ELISA? In quantitative ELISA, how much the product “colours” is corresponding to the amount of enzyme-linked antibody involved .

Smad proteins in the resting state realize passive nucleo cytoplasmic shuttling, that is controlled by two opposing signals: the nuclear localization signal .

There are two ways for monoclonal antibody’s growth: one is to place them in laboratory flasks (In Vitro), and the other is in the stomach lining of mice (fa.

An antibody elsa test works when a blood sample and the conjugate specific for that test are combined together. The conjugate contains an antibody that has a.

Coli strand. Transformation is the process in which there is a genetic alteration in a cell from uptaking genetic material from the environment. This can aff.

The Haloperidol will be administered at varying concentrations and exposure times. Using quantitative polymerase chain reaction (qPCR) will monitor the ampli.

The first signal is implemented by microbial molecules and cytokines that that bind to TLR receptor and induces the up-regulation the transcription of nuclea.

pneumoniae does not produce proteases and cannot break down proteins via proteolysis. The fat hydrolysis test was performed to determine whether our bacteria.

The sulfa medications, for example, sulfonamides hinder a basic compound - dihydropteroate synthase- - in this procedure. Once the procedure is halted, the m.


Cell Structure

The first life forms on Earth existed as single cells which were essentially a string of DNA enclosed in a cell wall, not dissimilar to modern bacterial cells (prokaryotes). After millions of years, multicellular organisms started to form containing organelles which specific functions. In fact, scientists believe that organelles such as mitochondria and chloroplasts arose when eukaryotic cells engulfed a bacteria. The bacteria remained in the cell, providing the organism with a handy supply of energy.

Microscopy

The microscope was first invented over four hundred years ago and it revolutionised science. For the first time scientists were able to visualise and confirm the existence of bacteria, laying the groundwork for the germ theory of disease which saved millions of lives. The first ever microscope was essentially two lenses inside a tube which resembled an empty loo roll but since then microscopes have come a long way with incredible powers of magnification.

The light microscope is the type of microscope you’ll have used in school. It uses light to magnify objects up to 1,500x their actual size. They have a resolution of approximately 0.2 μm which isn’t large enough to visualise any of the smaller organelles, such as ribosomes and lysosomes. They are more commonly used for visualising whole cells or tissues. An advantage of light microscopy is that it can visualise living cells so we can watch behaviours such as cell division in real time.

Laser-scanning confocal microscopes are an advanced type of light microscope which use an intense beam of light (a laser) to scan samples tagged with a fluorescent dye. When the laser hits the dye, the dye emits light which can be used to form an image.

Confocal microscopy image of a carnivorous plant. Credit: Dr. Igor Siwanowicz

The transmission electron microscope (TEM) is more powerful than a light microscope and has a high enough resolution (around 0.0002 μm) to visualise individual organelles. A TEM uses electromagnets to focus a beam of electrons at a sample. Electrons have a much shorter wavelength compared to visible light which means higher-resolution, detailed images can be produced. A disadvantage of TEM is that the sample needs to be fixed and placed in a vacuum, which means that live cells cannot be used.

The scanning electron microscope (SEM) has a lower resolution (around 0.002 μm) than the TEM but they can produce 3D images of cells and organelles. They emit a beam of electrons towards a sample, knocking electrons off it which are used to build an image. Like TEMs, SEMs cannot be used with live cells. Both types of electron microscope are pretty big and expensive so you’ll only find them in specialised research facilities and hospitals.

Light microscope TEM SEM
Maximum magnification 1 500 x 1 000 000 x 500 000 x
Maximum resolution 0.2 μm 0.0002 μm 0.002

Preparation of a microscope slide

Cut a really thin layer from your sample. If the specimen is too thick, light will not be able to pass through it.

Place a drop of water onto the microscope slide.

Using tweezers, place your specimen onto the drop of water and place a coverslip on top.

Add a stain if needed - place a droplet next to the coverslip and allow it to absorb across the sample.

Calibrating the eyepiece graticule and the stage micrometer

If you wanted to measure the size of your specimen, you’ll first need to align the eyepiece graticule and the stage micrometer which are little rulers which are found on the lens and the stage respectively. To do the calibration you need to carry out the following steps:

Place the stage micrometer on the stage and focus the lens so that you can clearly see the divisions.

Align the eyepiece graticule with the stage micrometer.

Each division of the stage micrometer is 0.1 mm. If the eyepiece graticule spans a total of three divisions, then we know that the total length of the eyepiece graticule is 0.2 mm.

The graticule is divided by a scale from 0 to 100, which means that each individual division is a length of 0.002 mm.

Now we can take away the stage micrometer and add our sample, using the eyepiece graticule to measure its size.

The nuclei of these cheek cells can be easily identified after staining with methylene blue. Credit: microscopemaster

Staining

Most biological samples are transparent and need to be stained to increase the contrast between different organelles so that they can be easily seen under the microscope. Different stains are used for different organelles: methylene blue is used to visualise DNA whereas eosin stains the cytoplasm. Iodine is often used for staining plant tissues.

Magnification

Make sure you understand the difference between magnification and resolution. Magnification is how enlarged the image is compared to the original object. Resolution is defined as how well a microscope distinguishes between two points that are close together (i.e. how much detail it can make out). Light microscopes have a much lower resolution, so produce less detailed images, compared to electron microscopes.

You can work out the magnification of a specimen viewed under a microscope using the equation:

Let’s say we magnify a 2 μm bacterial cell to form an image which is 16 cm long. The magnification we must have used is:

Convert both into the same units. 16 cm = 160 mm = 160,000 μm

160,000 / 2 = 80,000 x magnification

Ultrastructure of eukaryotic cells

All organisms are divided into two different domains: eukaryotes and prokaryotes. More complex, multicellular organisms are classed as eukaryotes whereas single-celled bacteria are prokaryotes. Eukaryotic cells, such as the cells of animals, plants and fungi may contain the following organelles:

Nucleus - contains DNA which controls the activities of the cell by containing the base sequences (the ‘instructions’ needed to make proteins. The DNA is associated with histone proteins and referred to as chromatin which is wound into structures called chromosomes.

Nucleolus - this is a region within the nucleus where ribosomes are made.

Nuclear envelope - a double membrane which surrounds the nucleus. It contains pores which allows small molecules (like single stranded RNA) to pass into the cytoplasm but keeps hefty chromosomes safely inside its walls.

Rough endoplasmic reticulum (RER) - the RER is an extension of the nuclear envelope and is coated with ribosomes. It facilitates protein synthesis by providing a large surface area for ribosomes. It then transports the newly synthesised proteins to the Golgi apparatus for modification.

Smooth endoplasmic reticulum (SER) - synthesises lipids including cholesterol and steroid hormones (such as estrogen).

Golgi apparatus - made up of a group of fluid-filled membrane-bound flattened sacs surrounded by vesicles. It receives proteins from the RER and lipids from the SER. It modifies the proteins and lipids and repackages them into vesicles. The Golgi apparatus is also the site of lysosome synthesis.

Ribosomes - ribosomes are responsible for the translation of RNA into protein (protein synthesis). They either float freely in the cytoplasm or are stuck onto the rough endoplasmic reticulum.

Mitochondria - site of ATP production during aerobic respiration. It is self-replicating so can become numerous in cells with high energy requirements. It contains a double membrane with folds called cristae, which provides a large surface area for respiration.

Lysosomes - phospholipid rings which contain digestive enzymes separate from the rest of the cytoplasm. Lysosomes engulf and destroy old organelles or foreign material.

Chloroplasts - the site of photosynthesis. It is enclosed by a double membrane and has internal thylakoid membranes arranged in stacks to form grana linked by lamellae. These structures are found only in plants and certain types of photosynthesising bacteria or protoctists.

Plasma membrane - consists of a phospholipid bilayer with additional proteins to serve as carriers. It also contains cholesterol to regulate membrane fluidity. The plasma membrane contains the cell contents and holds the cell together, whilst controlling the movement of substances into and out of the cell.

Centrioles - these are bundles of microtubules which form spindle fibres during mitosis in order to pull sister chromatids apart. They are also important for the formation of cilia and flagella. They are not found in plant and bacterial cells.

Cell wall - a rigid structure made of cellulose (in plants), chitin (in fungi) and murein (in prokaryotes) which provides support to the cell.

Flagella - a tail-like structure which are made up of bundles of microtubules. The microtubules contract to make the flagellum move and propel the cell forward. Cells with a flagellum include sperm cells, which use it to swim up the fallopian tubes to fertilise the egg cell.

Cilia - finger-like projections found on the surface of some cells. These also contain bundles of microtubules which contract to make the cilia move. Cilia are found on epithelial cells lining the trachea and move to sweep mucus up the windpipe.

Middle lamella - the middle lamella is found outside of the cell walls in plant cells. It is responsible for sticking plant cells together and providing stability. It is mostly made of a substance called calcium pectate.

Amyoplasts - these are plant storage granules which contain starch and are mostly found in bulbs and tubers. Amyoplasts can convert the starch back into glucose when the plant cell needs more glucose for respiration.

Vacuole - the vacuole is an organelle which stores cell sap and may also store nutrients and proteins. It helps to keep plant cells turgid. Some vacuoles can perform a similar function to lysosomes and digest large molecules.

Tonoplast - the tonoplast is a membrane which surrounds the vacuole which functions to separate the vacuole from the rest of the cell.

Plasmodesmata - plasmodesmata are narrow channels of cytoplasm within the cell walls of plants. It allows two neighbouring plant cells to transport substances between them and to communicate.

Pits - pits are regions of a plant cell where the cell wall becomes very thin. Pits are arranged in pairs so that the pit of one plant cell is aligned with the pit of another plant cell. Like plasmodesmata, pits allow neighbouring plant cells to exchange substances.

Protein production

During the production of proteins, different organelles function like an assembly line in a factory, each tweeking the protein a little until it ends up neatly packaged in a vesicle and released from the factory floor and ready to do its job in the cell. Proteins are first made on ribosomes which are either floating alone in the cytoplasm or attached to the rough ER. The long polypeptide chain is folded at the rough ER and transported to the Golgi apparatus inside vesicles. At the Golgi, they are modified and processed by various enzymes. The protein may have a carbohydrate chain stuck onto its surface, or the addition of a sulfate or phosphate group. The dolled up protein is then placed into another vesicle which travels to the part of the cell where the protein is needed. If the protein is a carrier protein, the vesicle will deliver the protein to the plasma membrane where it will be incorporated.

Cytoskeleton

Cells contain an important structure within its cytoplasm called the cytoskeleton. It’s a miniature skeleton that gives the cell shape and keeps organelles in position. It is made up of small tubes of protein called microtubules which form a network throughout the cell. The main functions of the cytoskeleton are:

Provides mechanical strength to cells

Allows the movement of organelles within the cell

Enables movement of the cell

Comparing eukaryotic and prokaryotic cells

Prokaryotes and eukaryotes share some of the same organelles (cytoplasm, cell membrane, ribosomes) but there are some important differences:

  • Prokaryotes have no membrane-bound organelles (so no mitochondria, Golgi, endoplasmic reticulum, nucleus etc). Their DNA floats freely in the cytoplasm.
  • Their DNA consists of a single circular chromosome whereas DNA in eukaryotes is linear and wrapped around chromosomes.
  • Prokaryotes have extra bits of DNA in the form of small circular plasmids.
  • Prokaryotes have smaller ribosomes (70S) compared to eukaryotic ribosomes (80S).
  • Eukaryotes like plants and fungi have cell walls made of cellulose and chitin. Bacterial cell walls are made of murein (a type of glycoprotein).
  • Prokaryotic cells are much smaller than eukaryotic cells.
  • Both prokaryotes and eukaryotes can have flagella but those found in prokaryotes are made of a protein called flagellin whereas in eukaryotes they are formed from microtubules.

Prokaryotes have some organelles that are absent from eukaryotic cells. These include:

Pili - pili are hair-like structures which stick out from the plasma membrane. They are used to communicate with other cells (including the transfer of plasmids between bacteria).

Mesosomes - the mesosome is a folded portion of the inner membrane. While some scientists believe that it plays a role in chemical reactions, such as respiration, other scientists doubt whether it even exists and think that it may just be an artefact produced during the preparation of bacterial samples for microscopy.

Plasmids - plasmids are small, circular rings of DNA which are separate from the main chromosome. They house genes which are not crucial for survival but might prove useful - such as antibiotic-resistance genes, for example. Plasmids can replicate independently from the main chromosomal DNA.

Slime capsule - in addition to a cell wall, some bacteria also have a capsule which is made of slime. The main function of the capsule is to protect the bacterium against an immune system attack.

Cell fractionation

Cell fractionation is a technique which separates organelles according to their density - you might want to do this if you want to visualise certain organelles under the microscope separately. It involves bursting the cell surface membrane to release the organelles and spinning the cell solution at really high speeds.

Homogenisation - the first step of cell fractionation is homogenisation. This is where you break apart the plasma membrane to release the organelles. This can be done by vibrating the cells or by breaking them apart in a blender. It is important that this cells are placed into a solution which is ice-cold, isotonic and buffered.

Ice-cold - the solution needs to be ice-cold to slow down the activity of enzymes. This is important because some enzymes will degrade organelles (such as the enzymes found inside lysosomes) so we need to reduce their activity to preserve the cell’s organelles.

Isotonic - the solute concentration (and therefore water potential) of the solution needs to be the same as the cells that have been broken down, otherwise water would move into the organelles by osmosis, resulting in damage

Buffered - adding a buffer to a solution ensures the pH stays constant. This is important because proteins are denatured by changes in pH - remember that proteins are a key component of various organelles.

Filtration - the homogenised solution is filtered to remove any tissue debris. The organelles are small enough to pass through the holes of the filter paper so will be present in the filtrate.

Ultracentrifugation - this is where we spin the filtrate at increasing speeds. The heaviest organelles will sink to the bottom of the test-tube, forming a pellet. We can transfer the remaining solution (the supernatant) to a separate test tube, which will be spun at a slightly higher speed. This is repeated until you obtain the organelle that you want. Remember that the organelles will be separated from the solution from the heaviest to the lightest. Nuclei will come out of the solution first, followed by mitochondria, then lysosomes, then the endoplasmic reticulum. Ribosomes will be the last organelles to form a pellet, since these are the lightest organelles in a cell.


Peer review information Nature Protocols thanks Gary Laevsky, Timo Zimmermann and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

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Results

Identification of CUP-2 and mutant phenotypes

The cup-2 gene was identified on the basis of the ar506 mutation that results in decreased endocytosis in scavenger cells called coelomocytes in Caenorhabditis elegans (Fares and Greenwald, 2001a). cup-2(ar506) worms have a temperature-sensitive defect in endocytosis by coelomocytes (Fig. 1A,B). pmyo-3::ssGFP transgenic worms express GFP fused to a signal sequence and expressed in body wall muscles: this GFP is secreted into the body cavity and is endocytosed, and subsequently degraded, primarily by the coelomocytes (Fares and Greenwald, 2001a). Mutations that decrease endocytosis by coelomocytes result in the accumulation of GFP in the body cavity and a decrease in the size of the GFP-filled compartments in coelomocytes (Dang et al., 2004 Fares and Greenwald, 2001a Patton et al., 2005). In addition to this temperature-sensitive endocytosis defect, cup-2(ar506) worms are sick or nonviable at 25°C, such that 90-95% of L4 larvae shifted from 20°C to 25°C die the rest of the worms survive and lay eggs, most of which are nonviable.

We determined that CUP-2 corresponds to open reading frame F25D7.1 based on phenocopy by RNA-mediated interference (RNAi), sequence analysis of the ar506 allele and transgenic rescue of the mutant phenotypes (Fig. 1C supplementary material Fig. S1A Fig. 2). The ar506 allele is a nucleotide change in the first exon that results in an early stop codon and therefore represents a predicted null mutant of cup-2 (Fig. 1C supplementary material Fig. S1B). CUP-2 is one of two C. elegans Derlin proteins and shows the highest identity to human Derlin-1 (Fig. 1D supplementary material Fig. S1B). Derlin proteins are conserved proteins that function in ERAD and have four predicted transmembrane domains, with both the N- and the C-termini being cytoplasmic (Fig. 1D, supplementary material Fig. S1B) (Hitt and Wolf, 2004 Knop et al., 1996 Lilley and Ploegh, 2004 Lilley and Ploegh, 2005 Ye et al., 2004). Consistent with this basic cell biological function, fusion of GFP to the CUP-2 promoter resulted in ubiquitous expression of GFP in all tissues, including coelomocytes (Fig. 1E).

We confirmed that CUP-2 functions in ERAD based on two criteria. First, cup-2(ar506) was synthetically lethal at 20°C with a null mutation in ire-1, the worm orthologue of the UPR sensor IRE1 (Fig. 1F) (Shen et al., 2001). Similarly, a yeast Δder1 Δire1 strain was nonviable at 37°C (Hitt and Wolf, 2004). cup-2 pek-1 and cup-2 atf-6 double mutants only showed slight increases in lethality (Fig. 1F). Second, in the cup-2(ar506) mutant, or after reducing CUP-2 levels by RNAi, the UPR was activated in most cells, including coelomocytes, as determined by the induction of GFP expression from an hsp-4::GFP reporter (Fig. 1G,H supplementary material Fig. S1A). A similar effect of cup-2 RNAi on induction of hsp-4::GFP expression in intestinal cells has been reported previously (Ye et al., 2004). Unlike the endocytosis defect, the UPR induction due to cup-2(ar506) is not temperature sensitive. Furthermore, the activation of the UPR response is absent in an xbp-1 null mutant, indicating that it is dependent on the site-specific cleavage and activation of xbp-1 mRNA by IRE-1 (supplementary material Fig. S1A) (Calfon et al., 2002).

RNAi of R151.6, the second worm Derlin protein, did not result in detectable UPR activation in any cells of wild-type animals it also did not affect endocytosis by coelomocytes (supplementary material Fig. S1A). However, CUP-2 and R151.6 exhibited partially redundant function in activating the UPR, because RNAi of R151.6 in cup-2(ar506) resulted in a dramatic increase in XBP-1-dependent hsp-4::GFP expression (supplementary material Fig. S1A).

Rescue of endocytosis and ERAD defects by CUP-2 homologues. (A) Quantification of the sizes of the GFP-filled vesicles in coelomocytes of cup-2(ar506) pmyo-3::ssGFP strains shown in supplementary material Fig. S2. 1 pixel=∼0.001 μm 2 . (B) Quantification of the average intensity per pixel of GFP in the nuclei of coelomocytes of the cup-2(ar506) hsp-4::GFP strains shown in supplementary material Fig. S2.

Rescue of endocytosis and ERAD defects by CUP-2 homologues. (A) Quantification of the sizes of the GFP-filled vesicles in coelomocytes of cup-2(ar506) pmyo-3::ssGFP strains shown in supplementary material Fig. S2. 1 pixel=∼0.001 μm 2 . (B) Quantification of the average intensity per pixel of GFP in the nuclei of coelomocytes of the cup-2(ar506) hsp-4::GFP strains shown in supplementary material Fig. S2.

Rescue of UPR activation and endocytosis defects by Derlin proteins

To determine whether CUP-2 function is conserved, we expressed several Derlin proteins from other species in cup-2 mutant worms. The CUP-2 promoter drove Derlin homologue expression in these transgenic animals. We observed essentially two patterns of rescue (Fig. 2 supplementary material Fig. S2). C. elegans CUP-2 and R151.6, and human Derlin-1 and Derlin-3 rescued both endocytosis and UPR activation phenotypes. These results indicate that both activities of CUP-2 are also found in mammalian Derlin proteins. Yeast Der1p and Dfm1p, and human Derlin-2 did not rescue either defect. Furthermore, yeast Dfm1p exacerbated the UPR activation, but not the endocytosis defect, suggesting that it might function as a dominant-negative protein that interferes with R151.6 activity in the ER.

Analysis of MCA-3 levels at the plasma membrane

To determine whether the loss of CUP-2 also affects the endocytic trafficking of transmembrane proteins, we analyzed the levels of the Ca 2+ pump CUP-7/MCA-3. MCA-3 was originally identified on the basis of mutations that disrupt endocytosis (Fares and Greenwald, 2001a). A functional GFP::MCA-3 fusion localizes to the plasma membrane (supplementary material Fig. S3A) (Bednarek et al., 2007). GFP::MCA-3 does not localize to the large vacuoles in a cup-5 mutant that is defective in lysosome-mediated degradation, indicating that it is not normally transported to the lysosome before or after endocytosis (supplementary material Fig. S3A) (Fares and Greenwald, 2001b Treusch et al., 2004). By contrast, GFP::MCA-3 accumulates to the membranes of expanded mCherry::RAB-11-positive recycling endosomes in an rme-1 mutant, indicating that MCA-3 continuously cycles between the plasma membrane and endosomes (supplementary material Fig. S3A,B) (Grant et al., 2001 Lin et al., 2001). RME-1 is an EH-domain-containing protein that is required for the exit from recycling endosomes loss of RME-1 results in expanded recycling endosomes (Grant et al., 2001 Lin et al., 2001).

cup-2(ar506) (or cup-2 RNAi) results in increased levels of GFP::MCA-3 at the plasma membrane as measured by quantitative microscopy (Fig. 3A,B and data not shown). Western blot analysis of immunoprecipitated GFP::MCA-3 indicated that the increased GFP::MCA-3 signal is due to increased levels of GFP::MCA-3 protein in cup-2 mutant worms (Fig. 3C,D). From three independent immunoprecipitation experiments, there was a 1.73±0.06 increase in the levels of GFP::MCA-3 in the coelomocytes of cup-2 mutant worms relative to the wild type this is in agreement with the measurements from microscopy (Fig. 3A,B). By contrast, GFP::MCA-3 does not accumulate at the plasma membrane of cup-5 mutants with strongly reduced lysosome function nor in a cup-4 mutant, which displays an even more severe general endocytosis defect than the cup-2 mutant (supplementary material Fig. S3A,C) (Patton et al., 2005 Treusch et al., 2004). Neither cup-2 nor cup-4 mutations completely block endocytosis in coelomocytes. This indicates that the accumulation of GFP::MCA-3 at the plasma membrane is not a general consequence of decreasing internalization rates. Furthermore the increase of GFP::MCA-3 at the plasma membrane is probably independent of the UPR activation defect of cup-2(ar506) because this increase is evident at 20°C and not at 15°C, which correlates with the endocytosis phenotype and not the UPR activation defect (Fig. 3A,B). Finally, we checked the levels of two other integral membrane proteins that are expressed under the control of the same coelomocyte promoter used to express GFP::MCA-3. Mannosidase II::GFP localizes to the Golgi complex whereas GFP::CUP-5 localizes to lysosomes in coelomocytes (Treusch et al., 2004). Neither protein showed increased levels in cup-2 mutants, indicating that the increase in MCA-3 levels is not due to an increase in general secretion or promoter activity (supplementary material Fig. S4A-D).

Analysis of endocytic markers. (A) Confocal micrographs of coelomocytes in adult hermaphrodites of the indicated genotypes, grown at 15°C or 20°C, and expressing GFP fusions to MCA-3. (B) Quantification of the plasma membrane fluorescence of the cells shown in A. (C) Western blot probed with anti-GFP antibodies of the indicated strains carrying the same GFP::MCA-3 transgene. The indicated bands are GFP::MCA-3 (197 kDa) and GFP (27 kDa). The GFP::MCA-3 transgene also includes DNA that expresses GFP in the pharynx of worms. The fluorescence intensity of this pharyngeal GFP was identical in the different strains. (D) Confocal micrographs of coelomocytes in wild type (WT) or cup-2(ar506) (cup-2) adult hermaphrodites grown at 20°C and expressing GFP fusions to the indicated proteins. CHC, clathrin heavy chain sER, cytochrome b5 Golgi, mannosidase II. (E) Western blot of proteins isolated from wild-type and cup-2(ar506) worms expressing clathrin heavy chain fused to GFP and probed with antibodies against GFP, RME-1 or actin. Equal amounts of protein were loaded in each lane. (F) Confocal images of coelomocytes at the indicated times after BSA-Rhodamine was injected into the body cavities of worms. Arrowheads indicate BSA-Rhodamine localized to lysosomes. Red, BSA-Rhodamine, Green, RME-8::GFP or GFP::CUP-5. Scale bars: 5 μm.

Analysis of endocytic markers. (A) Confocal micrographs of coelomocytes in adult hermaphrodites of the indicated genotypes, grown at 15°C or 20°C, and expressing GFP fusions to MCA-3. (B) Quantification of the plasma membrane fluorescence of the cells shown in A. (C) Western blot probed with anti-GFP antibodies of the indicated strains carrying the same GFP::MCA-3 transgene. The indicated bands are GFP::MCA-3 (197 kDa) and GFP (27 kDa). The GFP::MCA-3 transgene also includes DNA that expresses GFP in the pharynx of worms. The fluorescence intensity of this pharyngeal GFP was identical in the different strains. (D) Confocal micrographs of coelomocytes in wild type (WT) or cup-2(ar506) (cup-2) adult hermaphrodites grown at 20°C and expressing GFP fusions to the indicated proteins. CHC, clathrin heavy chain sER, cytochrome b5 Golgi, mannosidase II. (E) Western blot of proteins isolated from wild-type and cup-2(ar506) worms expressing clathrin heavy chain fused to GFP and probed with antibodies against GFP, RME-1 or actin. Equal amounts of protein were loaded in each lane. (F) Confocal images of coelomocytes at the indicated times after BSA-Rhodamine was injected into the body cavities of worms. Arrowheads indicate BSA-Rhodamine localized to lysosomes. Red, BSA-Rhodamine, Green, RME-8::GFP or GFP::CUP-5. Scale bars: 5 μm.

Therefore, at least one transmembrane protein, MCA-3, accumulates at the surface of coelomocytes in the absence of CUP-2. Given this result and the endocytosis defect, we assayed other markers of the endocytic pathway in coelomocytes.

Analysis of endocytosis markers in cup-2 mutant coelomocytes

To better understand the nature of the endocytosis defect in cup-2 mutants we analyzed the morphologies of all major membrane bound organelles of the endocytic and secretory pathways in cup-2(ar506) coelomocytes using a set of GFP-tagged markers that we developed (Fig. 3D). There was no change in the localization of GFP-RME-1, a recycling endosome and plasma-membrane-localized EH-domain-containing protein required for endocytic traffic in coelomocytes (Fares and Greenwald, 2001a Grant et al., 2001). By contrast, there was a significant increase in the levels of GFP-tagged clathrin heavy chain (CHC) at the surface of cup-2(ar506) coelomocytes (Greener et al., 2001). This increase in membrane CHC localization was not accompanied by a change in the absolute levels of CHC in cup-2(ar506) worms, indicating that the increased clathrin signal probably indicates increased clathrin assembly on membranes (Fig. 3E).

RAB-7, a marker for late endosomes, and CUP-5, a marker for lysosomes, both showed normal staining patterns, albeit of compartments that were reduced in size this reduction in the size of endosomes or lysosomes is a general defect seen in mutations that block coelomocyte endocytosis (Grant and Hirsh, 1999 Patton et al., 2005 Treusch et al., 2004) (Fig. 3D). There was an elaboration and a dispersal of the ER in cup-2(ar506) coelomocytes (Fig. 3D). Both of these phenotypes are seen during hyperactivation of UPR in mammalian plasma cells and in other mutations that block coelomocyte endocytosis (Patton et al., 2005 Shaffer et al., 2004 Sriburi et al., 2004). Finally, there was no obvious change in the localization of mannosidase II, a transmembrane Golgi marker (Fig. 3D) (Patton et al., 2005 Rolls et al., 2002).

We also determined whether the loss of CUP-2 results in a delay in transport of endocytosed BSA-Rhodamine to lysosomes. This experiment was feasible because although the rate of endocytosis of soluble molecules was reduced in the cup-2 mutant, internalization was not blocked. Wild-type and cup-2 mutant worms that express the endosomal marker RME-8::GFP or the lysosomal marker GFP::CUP-5 were injected with BSA-Rhodamine in their pseudocoeloms. In wild-type and cup-2 mutant worms, all of the BSA-Rhodamine was found in RME-8-positive, CUP-5-negative endosomes after 5 minutes of uptake, as has been previously shown (Dang et al., 2004 Treusch et al., 2004). In both wild-type and cup-2 mutant worms, we first detected BSA-Rhodamine in RME-8-negative, CUP-5-positive lysosomes 10 minutes into the time course (arrowheads in Fig. 3F). Therefore, the loss of CUP-2 does not affect the progress of endocytosed solutes and membrane through the endo-lysosomal pathway.

Therefore, the only defect that we detect in the absence of CUP-2, so far, is an accumulation of CHC at the surface of coelomocytes. This accumulation has not been observed in other mutants that disrupt coelomocyte endocytosis and is therefore not due to a general reduction in internalization (Bednarek et al., 2007 Grant et al., 2001 Patton et al., 2005 Sato et al., 2005 Xue et al., 2003 Zhang et al., 2001).

Given the roles of Derlin proteins in degrading misfolded proteins in the ER, we hypothesized that misfolded protein accumulation could explain the increased MCA-3 levels at the surface of cup-2 mutant coelomocytes. We therefore did studies in cultured cells where this idea could be tested.

Mammalian Derl1 RNAi phenotypes

The rescue of cup-2 mutant phenotypes by mammalian Derlin proteins suggests functional conservation. We therefore initiated studies in mammalian cells to allow for pulse-chase and biochemical manipulations that are not currently feasible in worms. We chose murine RAW264.7 macrophages because they are analogous to coelomocytes and therefore allow for a smooth transition for comparative analysis between the worm and the mammalian work.

We made a stable Derl1 RNAi clone in RAW264.7 cells. In these cells, Derlin-1 levels were 8.2±0.9% and Derlin-2 levels were 133.5±14.8% of that in the wild type (Fig. 4A). Derlin-3 was not detectable in the wild type or Derlin-1 RNAi clone (Fig. 4A). The increase in Derlin-2 levels is consistent with previous results showing that both Derlin-2 and Derlin-3 are upregulated by the UPR, which has probably been activated because of the reduced Derlin-1 levels (Oda et al., 2006).

Previous studies have shown that at least some of the low-density lipoprotein receptor (LDLR) at the plasma membrane is degraded by the proteasome (Martin de Llano et al., 2006 Miura et al., 1996). We therefore checked the levels of the LDLR at the plasma membrane by incubating live cells with antibodies at 4°C, followed by fixation and imaging. We detected a sevenfold increase in the levels of LDLR at the plasma membrane of the Derl1 RNAi clones relative to RAW264.7 cells or to Mcoln1 RNAi stable clones that were used as a control (Fig. 4B,C). Mcoln1 encodes mucolipin 1, which is an endosomal/lysosomal-localized protein required for efficient lysosomal trafficking and lysosomal degradation of endocytosed proteins in RAW264.7 cells (Thompson et al., 2007). We also checked the levels of Fcgamma receptors (FcR) using an antibody that recognizes CD16, CD32 and CD64 (Unkeless, 1979). Although the trafficking itinerary from the plasma membrane of these proteins is not clear, we also detected a fourfold increase in the levels of FcR at the plasma membrane of the Derl1 RNAi clone relative to RAW264.7 cells or to an Mcoln1 RNAi stable clone that was used as a control (Fig. 4B,C). Therefore, similarly to worm cup-2 mutants, reduced Derlin-1 levels result in an increase in the plasma membrane levels of at least two integral membrane proteins.

The increase in LDLR levels at the plasma membrane of the Derl1 RNAi clone did not result in a increase in LDL binding at the plasma membrane. We incubated cells with fluorescent LDL at 4°C before washing and fixing the cells at 4°C. We detected similar levels of LDL binding to the surfaces of RAW264.7, Derl1 RNAi, and Mcoln1 RNAi cells (Fig. 4D,E). This suggests that the substantial number of LDLRs at the plasma membrane of Derl1 RNAi cells that are unable to bind to LDL probably represent LDLR molecules whose extracellular ligand-binding domain is misfolded.

We wanted to directly assay the fates of transmembrane proteins at the plasma membrane that become misfolded. We first tested a high-salt and low-pH buffer (741 mM citric acid, 258.7 mM sodium citrate, pH 3.5) that is predicted to affect the conformation of the extracellular domains of many receptors. Treatment of cells with this misfolding buffer for 15 minutes at 4°C resulted in approximately one-fifth of the LDL binding to the cell surfaces relative to D-PBS-treated samples (Fig. 4D,E). This indicates that this buffer, although not lethal to cells (data not shown), causes conformational changes in the extracellular domains of receptors.

We then used this misfolding buffer to determine the parameters of the degradation of misfolded LDLR at the plasma membrane. We pre-incubated cells for 2 hours in cycloheximide to block translation and treated them with D-PBS (control) or with misfolding buffer. These cells were placed back in medium containing cycloheximide alone, with the proteasomal inhibitor MG132, or with the lysosomal inhibitor leupeptin. Samples were taken every 2 hours for western blot assays to measure the rates of disappearance of the plasma-membrane-localized mature form of the LDLR (confirmed using surface biotinylation assays, data not shown). There are three main conclusions from this analysis (Fig. 4F,G). First, induction of misfolding of surface LDLR results in a sharp increase in the rate of its degradation in normal cells (red vs black solid lines in Fig. 4G). Second, misfolded LDLR is degraded primarily by the proteasome (orange solid line in Fig. 4G) and is less dependent on lysosomal function (green solid line in Fig. 4G). This is consistent with previous studies of plasma-membrane-localized LDLR turnover in CHO cells under normal conditions, where a proteasomal pathway degrades 70% of these receptors and the rest are degraded in lysosomes (Martin de Llano et al., 2006). Third, reducing Derlin-1 levels stabilizes plasma membrane LDLR under normal conditions (black dashed lines in Fig. 4G) and after misfolding (red dashed lines in Fig. 4G). Therefore, misfolding of LDLR at the plasma membrane results in its rapid turnover in a proteasome and Derlin-1-dependent manner.

Loss of CUP-2 results in MCA-3 accumulation at the surfaces of cells. Reduced Derlin-1 levels results in an accumulation of misfolded LDLR at the plasma membrane and in a reduction in its rate of degradation. One possible explanation for this data is that some Derlin molecules localize to the plasma membrane and/or endosomes where they regulate the fates of misfolded proteins at the cell surface. We therefore assayed the localization of Derlin proteins in more detail.

Mammalian Derl1 RNAi phenotypes. (A) Western blots of proteins isolated from RAW264.7 cells or Derl1 shRNA-stable clones and probed with anti-Derlin-1, anti-Derlin-2, or anti-Derlin-3 antibodies. Anti-actin antibodies were used as loading control. (B) Confocal images of RAW264.7, Derl1 RNAi clones, or Mcoln1 RNAi clones stained with polyclonal antibodies that detect the extracellular domains of LDLR or Fcgamma receptors (FcR) and with FITC-labeled secondary antibodies. For each antibody, images were taken using the same exposure and magnification. (C) Quantification of the plasma membrane fluorescence staining of LDLR and FcR shown in B. (D) Confocal images of RAW264.7, Derl1 RNAi clones, or Mcoln1 RNAi clones stained with Bodipy FL-LDL after treating the cells with D-PBS or with a high-salt, low-pH misfolding buffer. All images were taken using the same exposure and magnification. (E) Quantification of the plasma membrane fluorescence staining of LDLR and FcR shown in B. (F) Representative western blots of the kinetics of degradation of mature LDLR. MF refers to treatment of cells with misfolding buffer. (G) Quantification of the mature LDLR levels over time from three kinetics of degradation experiments. RAW, RAW264.7 cells Derlin1KD, Derlin-1 shRNA clone MCOLN1KD, MCOLN1 shRNA clone PBS, D-PBS treatment MF, misfolding buffer treatment.

Mammalian Derl1 RNAi phenotypes. (A) Western blots of proteins isolated from RAW264.7 cells or Derl1 shRNA-stable clones and probed with anti-Derlin-1, anti-Derlin-2, or anti-Derlin-3 antibodies. Anti-actin antibodies were used as loading control. (B) Confocal images of RAW264.7, Derl1 RNAi clones, or Mcoln1 RNAi clones stained with polyclonal antibodies that detect the extracellular domains of LDLR or Fcgamma receptors (FcR) and with FITC-labeled secondary antibodies. For each antibody, images were taken using the same exposure and magnification. (C) Quantification of the plasma membrane fluorescence staining of LDLR and FcR shown in B. (D) Confocal images of RAW264.7, Derl1 RNAi clones, or Mcoln1 RNAi clones stained with Bodipy FL-LDL after treating the cells with D-PBS or with a high-salt, low-pH misfolding buffer. All images were taken using the same exposure and magnification. (E) Quantification of the plasma membrane fluorescence staining of LDLR and FcR shown in B. (F) Representative western blots of the kinetics of degradation of mature LDLR. MF refers to treatment of cells with misfolding buffer. (G) Quantification of the mature LDLR levels over time from three kinetics of degradation experiments. RAW, RAW264.7 cells Derlin1KD, Derlin-1 shRNA clone MCOLN1KD, MCOLN1 shRNA clone PBS, D-PBS treatment MF, misfolding buffer treatment.

Subcellular localization of CUP-2 and Derlin proteins

Der1p and Dfm1p in yeast, and Derlin-1 and Derlin-2 in humans, localize to the ER (Hitt and Wolf, 2004 Knop et al., 1996 Lilley and Ploegh, 2004 Lilley and Ploegh, 2005 Ye et al., 2004). To confirm the ER localization of CUP-2, we fused GFP to its C-terminus. Expression of this CUP-2::GFP fusion under the control of the coelomocyte promoter rescues both the endocytosis and the UPR activation defects of cup-2(ar506) in coelomocytes, indicating that the fusion protein is functional and that CUP-2 acts cell autonomously (Fig. 5A-C). In wild-type coelomocytes, CUP-2::GFP colocalizes extensively with the smooth ER marker cytochrome b5 and the rough ER marker TRAM, indicating that at steady state, the majority of CUP-2 molecules reside in the ER (Fig. 5D) (Rolls et al., 2002). However, in addition to the ER staining, we consistently detected CUP-2::GFP, but not the other ER markers, in peripheral organelles, suggesting that CUP-2 localizes to other compartments besides the ER (Fig. 5D, arrows). This peripheral staining was not of the Golgi complex that we visualized as discrete centralized puncta in coelomocytes (see Fig. 3D) (Bednarek et al., 2007 Dang et al., 2004 Patton et al., 2005 Treusch et al., 2004). At least a portion of the peripheral CUP-2 staining was endosomal because it appeared to colocalize with RAB-5, although the elaborate nature of the ER in coelomocytes precludes an unambiguous determination (Fig. 5D, arrows).

Subcellular localization of CUP-2. (A) Confocal images of pmyo-3::ssGFP or cup-2(ar506) pmyo-3::ssGFP adult hermaphrodites (left panels) and individual coelomocytes (right panels) expressing a CUP-2::GFP translational fusion from a coelomocyte-specific promoter. Worms were grown at 20°C. (B) Quantification of the sizes of the GFP-filled vesicles in coelomocytes of strains shown in A. 1 pixel=∼0.001 μm 2 . (C) Average intensity per pixel of GFP in the nuclei of coelomocytes of strains of the indicated genotypes expressing HSP-4::GFP. Worms were grown at 20°C. (D) Confocal images of coelomocytes co-expressing CUP-2::GFP and the smooth ER (sER) marker cytochrome b5 or CUP-2::GFP and the rough ER (rER) marker TRAM or CUP-2::GFP and the early endosome marker RAB-5, fused to mCherry. Arrows indicate structures that only contain CUP-2::GFP in the ER localization images and endosomes that contain both CUP-2 and RAB-5 in the bottom panels. Worms were grown at 20°C.

Subcellular localization of CUP-2. (A) Confocal images of pmyo-3::ssGFP or cup-2(ar506) pmyo-3::ssGFP adult hermaphrodites (left panels) and individual coelomocytes (right panels) expressing a CUP-2::GFP translational fusion from a coelomocyte-specific promoter. Worms were grown at 20°C. (B) Quantification of the sizes of the GFP-filled vesicles in coelomocytes of strains shown in A. 1 pixel=∼0.001 μm 2 . (C) Average intensity per pixel of GFP in the nuclei of coelomocytes of strains of the indicated genotypes expressing HSP-4::GFP. Worms were grown at 20°C. (D) Confocal images of coelomocytes co-expressing CUP-2::GFP and the smooth ER (sER) marker cytochrome b5 or CUP-2::GFP and the rough ER (rER) marker TRAM or CUP-2::GFP and the early endosome marker RAB-5, fused to mCherry. Arrows indicate structures that only contain CUP-2::GFP in the ER localization images and endosomes that contain both CUP-2 and RAB-5 in the bottom panels. Worms were grown at 20°C.

We determined the subcellular localization of endogenous mammalian Derlin proteins to confirm this extra-ER localization. We first used immunofluorescence comparing the localization of Derlin proteins to the ER-marker calreticulin. We observed significant colocalization of Derlin-1 with calreticulin and of Derlin-2 with calreticulin in RAW264.7 macrophages (Fig. 6A). However, there were vesicular structures that labeled for endogenous Derlin proteins but that did not contain calreticulin (arrows in Fig. 6A). We then transfected RAW264.7 cells with GFP- or YFP-tagged Rab5 (early endosomes), Rab7 (late endosomes) and Rab11a (recycling endosomes), that had been fixed, and immunostained to detect endogenous Derlin proteins and GFP or YFP. We saw significant colocalization between Derlin proteins and Rab5 and Rab7, indicating Derlin proteins are also found on endosomes (arrows in Fig. 6B). The Derlin proteins and Rab11a showed very limited, if any, colocalization in the perinuclear region (Fig. 6B). The colocalization of Derlin proteins with the Rab proteins is not an indirect consequence of Rab overexpression affecting ER integrity because we did not observe any colocalization between the Rab proteins and calreticulin in the same Rab-transfected cells (supplementary material Fig. S5).

We used immuno-electron microscopy to determine Derlin protein localization at a higher resolution. We first allowed cells to endocytose BSA-gold (15 nm) for 10 minutes to unambiguously label endosomal compartments (Fig. 6C). We then added anti-Derlin-1, anti-Derlin-2, or PBS (control) to cells before visualization using 6-nm-gold-conjugated secondary antibodies. RAW264.7 endosomes containing the 15-nm-gold particles showed peripheral staining for Derlin-1 (25/27 endosomes) and for Derlin-2 (24/25 endosomes). Only 1 out of 16 endosomes had 6-nm-gold particles in the control. Similar staining of the Derl1 RNAi cells revealed reduced staining for Derlin-1 (6/17 endosomes) but not for Derlin-2 (14/14 endosomes) (data not shown). Although we detected specific Derlin-1 and Derlin-2 labeling near the plasma membrane, we could not ascertain whether these represented actual plasma membrane labeling or labeling of subcortical organellar (ER, endosomes) membranes.

Finally, we fractionated the post-nuclear membrane fraction of murine RAW264.7 macrophages on a continuous iodixanol gradient to biochemically probe the presence of Derlin proteins in compartments besides the ER. Endogenous Derlin-1 and Derlin-2 was separated in two pools (boxed in Fig. 6D). Pool I (fractions 9-16) included ER membranes. Pool II (fractions 2-5) did not include ER membrane but did include Golgi complex, endosomes, and plasma membrane. Derlin-3 was not expressed in these cells (see Fig. 4A). These results indicate that Derlin proteins have a conserved localization to endosomal compartments in addition to the ER.

Subcellular localization of mouse Derlin proteins. (A) Confocal images of RAW264.7 or Derl1 RNAi cells stained for Derlin proteins and calreticulin (CRT). Arrows indicate vesicular structures that only stain for Derlin proteins. (B) Confocal images of RAW264.7 cells transfected with GFP-Rab5 (canine), YFP-Rab7a (canine), or GFP-Rab11a (canine) and then methanol-fixed and stained for Derlin proteins and GFP or YFP. Arrows indicate colocalization. (C) Electron micrographs of RAW264.7 cells that have endocytosed BSA-15-nm-gold particles and that were immunostained to detect Derlin-1 or Derlin-2. Secondary antibodies were conjugated to 6-nm-gold particles. Arrows indicate 6-nm-gold particles on the peripheries of endosomes. Scale bar: ∼0.1 μm. (D) Western blots of the subcellular fractions of RAW264.7 membranes. Gradients were fractionated from the top (top=1).

Subcellular localization of mouse Derlin proteins. (A) Confocal images of RAW264.7 or Derl1 RNAi cells stained for Derlin proteins and calreticulin (CRT). Arrows indicate vesicular structures that only stain for Derlin proteins. (B) Confocal images of RAW264.7 cells transfected with GFP-Rab5 (canine), YFP-Rab7a (canine), or GFP-Rab11a (canine) and then methanol-fixed and stained for Derlin proteins and GFP or YFP. Arrows indicate colocalization. (C) Electron micrographs of RAW264.7 cells that have endocytosed BSA-15-nm-gold particles and that were immunostained to detect Derlin-1 or Derlin-2. Secondary antibodies were conjugated to 6-nm-gold particles. Arrows indicate 6-nm-gold particles on the peripheries of endosomes. Scale bar: ∼0.1 μm. (D) Western blots of the subcellular fractions of RAW264.7 membranes. Gradients were fractionated from the top (top=1).


Discussion

Our studies can be summarized in the model shown in Fig 6E–G. Our previous study demonstrated the mutual inhibition between Ras activity and PI(3,4)P2 establishes polarity even in immobilized latrunculin A-treated cells in the presence of a chemoattractant gradient or caffeine (Fig 6E Arai et al, 2010 Wang et al, 2013 Li et al, 2018 ). We now propose an additional mechanism involving a specific vesicle recycling path which sharpens the back-to-front gradient in migrating Dictyostelium cells. PI(3,4)P2 disappears from protrusions at the leading edges of macropinocytic cups, and then accumulates on the macropinosomes at the end of the internalization process. The macropinosomes are processed into smaller satellite PI(3,4)P2-tagged vesicles which then dock at the back. The PI(3,4)P2 molecules incorporated at the back diffuse along the membrane toward the front, where they are degraded (Fig 6F). In more polarized cells, owing to a slower arrival time of PI(3,4)P2 vesicles and a slower decay rate of PI(3,4)P2 on the membrane, the PI(3,4)P2 signal at the back broadens and extends further to the front (Fig 6G). Our results suggest that a similar mechanism could exist in polarized migrating mammalian cells.

Excitable network hypothesis and the formation of macropinocytic cups

This model is consistent with the excitable network hypothesis, which has been proposed to explain the behaviors of migrating cells (Li et al, 2020 ). According to this hypothesis, propagating waves have an active region where front molecules are high and back molecules are low. This region is followed by a refractory region, where front molecules are very low and back components strongly accumulate (Xiong et al, 2010 Huang et al, 2013 Tang et al, 2014 Li et al, 2018 ). Here, our findings suggest macropinosomes are formed by a spreading wave whose trailing edge forms the base of the macropinocytic cups. This suggests that the base of the macropinocytic cups would be in a refractory state, which is heavily decorated by the back molecule PI(3,4)P2. If PI(3,4)P2 is delivered to the back of the cell as we propose, it would shut off this region and further polarize the cell.

Many other signaling molecules such as PTEN, Myosin II, IQGAP1 and 2, cortexillin I and II, RasGAP2 (RG2) and RapGAP3 (RG3) localize to the back of the cells (Ramalingam et al, 2015 Li et al, 2018 , 2020 ). Among those molecules, PTEN, RG2, RG3, and Myosin II are also reported to localize to the base of the cups or retracting protrusions. In addition, RG2 and RG3 bind to PI(3,4)P2 in vitro. Our results raise the possibility that the cells might be bringing the refractory state carrying many of those molecules from front to back.

Regulation of vesicle recycling controls polarity

In the reverse-fountain flow model, we were proposing that vesicular trafficking plays an important role in polarity. Yet, the vesicle recycling mechanism we described is more obvious in less polarized cells. How can we explain this apparent discrepancy? Highly polarized cells are generally characterized by their typical elongated morphology, which is a result of having a larger proportion of the cell with elevated backmarkers. In fact, our simulations showed that the relatively longer arrival time of the vesicles and the slower decay rate increased the back region. Subsequently, experiment measurements did show a slower decay rate in highly polarized cells. Thus, both experimental data and simulation results are consistent in supporting the reverse-fountain flow model (Fig 6 and Appendix Fig S6).

PI(3,4)P2 is a distinct signaling component during cell migration

In some studies of cell motility and cytoskeletal events, PI(3,4)P2 acts as a negative regulator (Lam et al, 2012 Ghosh et al, 2018 ). Huttenlocher’s group reported that Src-homology 2-containing inositol 5′ phosphatase (SHIP) limits the motility of neutrophils and their recruitment to wounds in live zebrafish (Yoo et al, 2010 ). Observations by Wu showed that PI(3,4)P2 defines the refractory period of the oscillation in cortical waves in mast cells (Xiong et al, 2016 Yang et al, 2017 ). Our lab reported that PI(3,4)P2 negatively regulates cell motility by inhibiting Ras activity, even in the absence of PIP3.

Our results depart from the canonical view that PI(3,4)P2 is only a byproduct of PIP3 hydrolysis (Czech, 2000 ). We demonstrated in mammalian neutrophils and in Dictyostelium cells that the level of PI(3,4)P2 was maintained on the plasma membrane under PI3K inhibition (Fig 3A–E). Additionally, in PI3K12Dictyostelium cells, PI(3,4)P2 still accumulated on the membrane at the back of the cells and the macropinosomes (Fig 3F–I). This suggests that a fraction of PI(3,4)P2 must come from a source other than PIP3 and there is regulation of enzymes that generate PI(3,4)P2 from PI3P or PI4P. As there are multiple phosphoinositide kinases in Dictyostelium that have not been characterized, it is possible that one of these corresponds to a novel PI3K.

Endocytic trafficking of PI(3,4)P2 as a universal means of establishing polarity

Our current findings reveal that the PI(3,4)P2 decorated macropinosomes and its connection with PI(3,4)P2 gradient at the rear of the plasma membrane in regulating polarity during cell migration. Consistently, there are reports suggesting recycling of PI(3,4)P2-tagged vesicles and other endocytic events play an important role in polarity of epithelial cells. A study by the Bryant group recently described the function of PI(3,4)P2 in apical domain morphogenesis in MDCK cysts. Apical PI(3,4)P2 is supplied by the endosomal pool, rather than conversion from basolateral PIP3 by SHIP1 and is a determinant of apical membrane identity (Roman-Fernandez et al, 2018 ). These observations reinforce the parallels between basolateral and apical surfaces of epithelial cells and front and back of migrating cells (Martin-Belmonte et al, 2007 Nelson, 2009 Bryant et al, 2014 ). Another parallel could be the establishment of polarity during cytokinesis, where PI(3,4)P2 is found elevated in the cleavage furrow (Li et al, 2020 ). The potential role of vesicular trafficking in cytokinesis has not been investigated.

Reverse-fountain flow model of PI(3,4)P2

During migration, the additional membrane required to enable the expending protrusions could come from unfolding of invaginations of the membrane or from vesicular trafficking.

Multiple different studies have suggested that cells migrate following a fountain flow model: Membrane precursor vesicles fuse with the anterior cell membrane at the protrusions, both the dorsal and ventral membranes flow toward the rear, and membrane is internalized at the rear (Lee et al, 1990 Tanaka et al, 2017 ). In support of the fountain flow models, it has been shown that blocking of vesicular trafficking is required for movement and that particles sticking on the outside of the cells as well as photobleached membrane patches flow from the front to the back.

In our reverse-fountain flow model, PI(3,4)P2-enriched vesicles are taken from macropinosomes at the anterior protrusion and eventually fuse to the membrane at the rear. Furthermore, plasma membrane PI(3,4)P2 at the rear diffuses toward the front. While this reverse-fountain flow model of PI(3,4)P2 sheds light on the role of this phospholipid in regulating polarity during cell migration, further study is needed to determine the direction of flow of other membrane components. One study showed that photoactivation of cAR1 did move toward the front, which support the reverse-fountain flow model (Traynor & Kay, 2007 ). In addition, vesicles containing adenylyl cyclase fuse with the back of Dictyostelium cells and vesicles carrying growth factor receptor fuse at the back of cells (Kriebel et al, 2008 Zoncu et al, 2009 ). A recent study by Moreau et al ( 2019 ) showed that migrating immature dendritic cells form macropinosomes at their leading edge which traverse the cytoplasm and ultimately release their fluid content at the back. This agrees with our model. Membrane folding, fountain flow model, or reverse-fountain flow models may operate in specific cells during different migratory modes.


Sample Preparation for Fluorescence Microscopy: An Introduction - Concepts and Tips for Better Fixed Sample Imaging Results

Multiple processing steps are required to prepare tissue culture cells for fluorescence microscopy. Experiments are generally classified as being either live or fixed cell microscopy. Fluorescence microscopy of live cells uses either genetically encoded fluorescent proteins (e.g. GFP, mcherry, YFP, RFP, etc.) or cell membrane-permeable, non-toxic fluorescent stains. Fluorescence microscopy of fixed cells uses a fixative agent that renders the cells dead, but maintains cellular structure, allowing the use of specific antibodies and dyes to investigate cell morphology and structure. Appropriate sample preparation is necessary to ensure high quality images are captured. Here we describe a number of concepts and considerations regarding the sample preparation process that can assist with automated digital fluorescence microscopy of fixed cells.

Cell Fixation

The goal of fixation is to maintain cellular structure as much as possible to that of the native or unfixed state during the processing steps and subsequent imaging. There are a number of fixation methods suitable for fluorescence microscopy that fall into two basic categories: aldehyde fixatives and alcohol fixatives. Organic solvents such as alcohols and acetone remove lipids and dehydrate the cells, while precipitating the proteins on the cellular architecture. Cross-linking aldehyde reagents form intermolecular bridges, normally through free amino groups, creating a network of linked antigens. Cross-linkers preserve cell structure better than organic solvents, but may reduce the antigenicity of some cell components, and require a permeabilization step to allow the antibody access to the specimen. Fixation with both methods may denature protein antigens, and for this reason, antibodies prepared against denatured proteins may be more useful for cell staining. As each method has its advantages and disadvantages and because different fixative methods can destroy or mask different epitopes, several different methods may need to be tested for optimal results.

Aldehyde fixatives cross link proteins and generally do an excellent job of preserving cell morphology. While they are generally slower acting than organic based fixatives, 4% formaldehyde for 10 minutes at room temperature is a good starting point. Glutaraldehyde will also cross link proteins, but results in significant autofluorescence and generally should be avoided or used in low concentration in conjunction with formaldehyde. Note that glutaraldehyde is the preferred fixative of choice for electron microscopy.

Formaldehyde vs. Paraformaldehyde vs. Formalin

Paraformaldehyde is a polymer of formaldehyde. It exists as a dry powder suitable for storage, and needs to be broken down into its monomeric component formaldehyde. This is accomplished by heating under basic conditions until it becomes solubilized. Commercial formaldehyde solutions generally referred to as formalin, often contain methanol to prevent re-polymerization into paraformaldehyde. A saturated formalin solution has formaldehyde content in water or buffer of 37- 40% and is sometimes referred to as &ldquo100% formalin&rdquo. Therefore using 10% formalin (1:10 dilution) is equal to using 3.7-4% formaldehyde for fixation. Since &ldquo100% formalin&rdquo contains up to 15% of methanol as a stabilizer, it can have a significant impact on fixation. This is mainly the result of membrane permeabilization by methanol, which results in the interference in the staining of membrane bound proteins.

Prepare a fresh 4% formaldehyde solution by dissolving paraformaldehyde powder into PBS (PH 7.4) using a stirring hot plate with the heater set to a medium setting until the liquid reaches approximately 60 °C. As the paraformaldehyde breaks down to formaldehyde it will dissolve. Once the solution is completely dissolved (approx 30 min), the solution is quickly chilled on ice back to room temperature prior to use. Typically, tissue culture cells are fixed for 10 minutes at room temperature with 4% paraformaldehyde in PBS followed by 2-3 washes with PBS to remove excess formaldehyde and stop the fixing reaction.

Organic solvents such as methanol rapidly precipitate proteins, maintaining structure when doing so. While the cellular protein cytoskeletal structure is maintained with methanol fixation, small molecules within will be lost during the subsequent processing steps because they are not precipitated. Because methanol precipitates proteins never use this method if genetically encoded fluorescent proteins are to be imaged or detected - activity from these proteins will be abolished with methanol fixation. Methanol is used cold (-20 °C) for 10-20 minutes. Using a combination of methanol and acetone (1:1) can sometimes improve results. Methanol is best for preserving structure while acetone improves permeabilization. Following fixation samples are washed with PBS 2-3 times to remove alcohol and rehydrate the specimen.

Another important considerationof a fixation protocol is the buffer selection. Ideally the buffer is isotonic in nature so as to not disrupt cellular structure, while maintaining a pH as close to physiologic as possible. The most frequently used buffer is phosphate buffered saline (PBS). When using aldehydes as fixatives avoid using amine-containing buffers, such as Tris, as they will react with the fixative.

Permeabilization

Fixation with the use of cross linking agents does not effectively diminish cell membrane structure. Therefore, formaldehyde fixation requires the cells to be permeabilized to allow antibodies access into the interior of cells. The large size and ionic nature of antibody proteins precludes them from gaining access without membrane disruption (Figure 1-2). Alcohol and acetone based fixation generally does not require this step.

Digitonin, Leucoperm and Saponin are relatively mild detergents that generate large enough pores for antibodies to pass through without completely dissolving the plasma membrane. These are suitable for antigens in the cytoplasm or the cytoplasmic face of the plasma membrane (Figure 1). Saponin acts by dissolving cholesterol present in the plasma membrane. When used at low concentrations, internal membranes remain intact so it is useful for labeling smaller molecules that exist in a soluble state within the cytoplasm. It should be prepared as a stock in DMSO, and is typically used at 0.5 to 1 mg/mL at room temperature for 10 to 30 minutes.

Triton X-100 is probably the most commonly used permeabilization agent for immunofluorescent staining. This detergent efficiently dissolves cellular membranes without disturbing protein-protein interactions (Figure 1). Triton is usually used at concentrations ranging from 0.1 to 1% for permeabilization. We typically use Triton X-100 or NP-40 at 0.1% in PBS at room temperature for 10 minutes. Besides cell membrane permeabilization, these detergents will partially dissolve the nuclear membrane making them suitable for nuclear antigen staining.

Figure 1. Antibody Accessibility with Unpermeabilized and Digitonin or Triton X-100 Permeabilized Cells. Antibodies can only access the exterior of aldehyde fixed unpermeabilized cells, while mild agents such as digitonin will permeabilize the plasma membrane allowing access to cytoplasmic interior, but not interior membrane bound organelles such as the nucleus of mitochondria. Stronger nonionic detergents, such as Triton X-100, permeabilize both the plasma membrane and interior membranes, allowing full access while still preserving cell structure.

SDS is an anionic detergent that is commonly used to denature proteins and provide them with a large negative charge for electrophoresis. It is useful as a permeabilizing agent to induce slight denaturation of fixed cells in order to reveal epitopes which may normally be masked from an antibody. It may extract small, poorly cross-linked proteins from fixed specimens, and should not be used on samples fixed by precipitation (e.g. methanol).

When using antibodies to stain cellular objects in specimens, it is necessary to &ldquoblock&rdquo the sample in order to reduce non-specific binding. Non-specific binding may occur for several reasons: unreacted fixative aldehydes may crosslink antibodies to inappropriate structures structures within the samples may &lsquotrap&rsquo antibodies or, if using polyclonal antibodies, low affinity IgG molecules may bind to inappropriate structures. These potential issues may be prevented by treating the specimens with a protein solution that will compete for non-specific binding sites prior to staining with antibodies. Commonly used blocking agents are bovine serum albumin (BSA), casein (or a solution of non-fat dry milk), gelatin, or normal serum obtained from the species of animal in which the secondary antibodies are made. Avoid using serum of the same species as the primary antibodies. Typically, the protein solutions are used at concentrations of 1 to 10% in buffer and the samples are treated after permeabilization for 10 to 30 minutes. It is also advisable to include a small amount of detergent (if the sample is to be permeabilized) to compete for hydrophobic interactions with the antibodies. In this case, low (around 0.1%) concentrations of Triton X-100 should be used, for example 3% BSA in PBS with 0.1% Triton X-100 or 5% fetal calf serum (FBS) in PBS with 0.1% Triton X-100 for 30 minutes at room temperature works well.

Antibodies for immunofluorescence are divided into two categories: primary and secondary antibodies. These two groups are classified based on whether they bind to antigens or proteins directly or target another (primary) antibody that, in turn, is bound to an antigen or protein. Primary antibodies can be labeled or unlabeled. Primary antibodies that have been covalently linked with a fluorescent moiety will have specificity and bind directly to the target as well as the tag necessary for imaging. This is known as direct immunofluorescence (Figure 2). By coupling the primary antibody with a fluorophore, direct immunofluorescence is faster than the indirect version because time-consuming washing and incubation steps are omitted. Thus, direct immunofluorescence is easier to handle and therefore suitable for the rapid analysis of samples in standardized experiments, for example in clinical practice. Unlabeled primary antibodies on the other hand will bind to the target but require a secondary antibody that will specifically recognize the antibody. This situation is called indirect immunofluorescence.

Figure 2. Directly labeled antibody fluorescence on unpermeabilized SK-BR-3 Cells. Anti-EGFR antibody covalently labeled with FITC and incubated with unpermeabilized SK-BR-3 cells counter stained with Hoechst 33342. Cells were imaged with a Cytation&trade 3 Cell Imaging Multi-Mode Reader (BioTek Instruments). Green fluorescence is located only on outer cell periphery as EGFR is a plasma membrane receptor.

Primary antibody binding to the specific antigen involves the Fab domain of the antibody, which in turn leaves the Fc domain exposed (Figure 3). The secondary antibody&rsquos Fab has been developed to recognize the Fc domain of the primary antibody. Since antibody&rsquos Fc domain is constant within the same animal class, only one type of secondary antibody is required to bind to many types of primary antibodies. This reduces the cost by labeling only one type of secondary antibody, rather than labeling various types of primary antibodies.


Figure 3. Structure of IgG Protein.

Primary antibodies can be either polyclonal or monoclonal in nature. Polyclonal antibodies are a heterogeneous mixture of antibodies directed against various epitopes of the same antigen. As they are generated from different B-cell clones they are immunochemically dissimilar and can have different specificities and affinities. While a number of different animal species, including goat, swine, guinea pig and cow, are routinely used to produce polyclonal antibodies, rabbits are most frequently used. Monoclonal antibodies are a homogeneous population of immunoglobulin directed against a single epitope. These antibodies are generated from a single B-cell clone isolated from a single animal and as such have the same Fab structure. While the vast majority of monoclonal antibodies are produced from mice, increasingly, they are being produced from rabbits as well.

Binding affinity and the resultant titer towards its antigen is different for every antibody. The use of monoclonal antibodies removes much of the variability, but batch to batch concentrations can vary even with the same monoclonal due to aggregation or denaturation. As a result the antibody concentration required and the incubation time used can vary lot to lot. Optimal assay development requires that the primary antibody be titrated for best results. Too little antibody results in an artificially low signal, due to a lack of target saturation. Too much antibody can result in nonspecific binding despite have used a blocking step. Because of the reagent cost, it is paramount that only as much antibody as required be used. However, for a one-off experiment, or a small series of experiments the time, effort and expense required for a full antibody titration is often not warranted. Typically, a dilution of 1:1000 for the commercially available monoclonal antibodies is used. Because of the low amounts of protein in the sample, it is best to use a buffer containing BSA to dilute the antibody, for example, PBS 0.1% Triton X-100 supplemented with 30 mg/mL of BSA. This can be made in bulk, filter sterilized and aliquoted. Use only enough antibody dilution to cover the sample, - for a typical well in a 96-well microplate this is 20-25 &muL. Most of the commercially available antibodies have sufficient titer to bind high copy number targets in 30-60 minutes at room temperature. Results with low copy number targets often are improved by overnight incubation at 4 °C, and shaking can help as well. An incubation time of 60 minutes at room temperature works well for most experiments, but an overnight incubation can be used as a convenient end of the day stopping point for sample processing even with high target number epitopes.

Figure 4. Direct vs. Indirect Antibody Labeling.

A secondary antibody is required with indirect immunofluorescence, where the primary antibody provides no means of detection and works by binding to a primary antibody, which directly binds to the target antigen. The use of secondary antibodies to indirectly detect target antigens requires additional process steps but can also offer significant advantages over labeled primary antibodies. Secondary antibodies increase the sensitivity through the signal amplification that occurs as multiple secondary antibodies bind to a single primary antibody (Figure 4). In addition, a given secondary antibody can be used with any primary antibody of the same isotype and target species, making it a more versatile reagent than individually labeled primary antibodies. This flexibility is a significant advantage of indirect immunofluorescence. Because the vast majority of primary antibodies are produced in just a few host animal species and most are of the IgG class, it is economical to produce and supply ready-to-use secondary antibodies for many methods and detection systems. From a relatively small number of secondary antibodies, many options are available for purity level, specificity and label type for a given application. A dilution of 1:500 for some commercially available secondary antibodies (Figure 5). As with primary antibodies, the low amounts of protein in the reagent makes it imperative that a buffer containing BSA or other non-specific protein be used to dilute the antibody, for example PBS 0.1% Triton X-100 supplemented with 30 mg/mL of BSA can be used for the dilution of secondary antibodies.

Figure 5. Fixed and three color fixed and stained HeLa cells. Mitochondria identified by a mouse anti-mitofilin primary antibody followed with a Rabbit antimouse IgG monoclonal antibody labeled with Texas Red (Red). Nuclei and actin filaments are identified by DAPI (blue) and AlexaFluor®-488-phalloidin (green) counterstaining respectively. Scale bar indicates 30 &mum.

It is important that the secondary antibody employed in the experiment is raised against the host species used to generate the primary antibody. Polyclonal primary antibodies are usually raised in rabbit, goat, sheep or donkey and are generally IgG isotypes. The secondary antibody therefore, will typically be an anti-IgG antibody. Monoclonal primary antibodies are commonly raised in mouse, rabbit and rat. For example, if the primary monoclonal antibody is a mouse IgG, you will need an anti-mouse IgG.

The use of pre-absorbed secondary antibodies reduces the risk of cross reactivity with other antibodies and should be used with multi-color experiments when several primary antibodies and their corresponding secondary antibodies are used simultaneously.

Pre-adsorption is an additional processing step where the secondary antibodies are passed through a column matrix containing immobilized serum proteins from potentially cross reactive agents. For example, only antibodies specific to rabbit IgG will pass through a pre-absorbing column containing immobilized mouse, human and horse IgG whereas, antibodies cross reacting will bind and stay adsorbed to the matrix.

Between each process step, there is usually a wash step to remove unbound excess reagents. The buffer used for these steps should take into account the buffer used for the process steps. As described previously, it is best to use a buffer system that is isotonic to preserve cellular structure and close to physiological pH. Washing, as the name implies is a series of aspiration and buffer reagent additions performed in sequence. Depending on the volume used, 2-3 cycles is sufficient for washing, with the final step being an aspiration to remove the fluid prior to the addition of the next processing reagent. PBS is commonly used, and a surfactant such as 0.1% Triton X-100 or 0.05% Tween&trade 20 can also be added. Note that it is important to make these steps in a timely fashion to prevent the sample from drying out.

Counter Stains

Counter stains are used for two different purposes reduction of background fluorescence or the identification of cellular organelles. Blue dyes, such as Evans blue can be used to reduce non-specific background fluorescence with fluorescein labeled samples. Because these dyes can mask weakly fluorescent desired binding they are not recommended for routine use. More importantly is the use of counter stains to identify cellular organelles and provide information regarding signal localization.

Figure 6. NIH 3T3 cell nuclei stained with DAPI.

The nucleus is the predominately targeted cellular organelle and can be identified through the use of stains that bind DNA (Figure 6). DAPI (diamidino-2-phenylindole) is a nuclear counter stain useful with multicolor fluorescent experiments. Its blue fluorescence differs significantly from fluorescein (green) or Texas red fluorescent probes used for other structures. It fluoresces brightly upon selectively binding to the minor groove of double stranded DNA. Its selectivity for DNA allows efficient staining of nuclei with little background from the cytoplasm. Other nuclear stains include Hoechst 33342, which is cell permanent and can be used with live as well as fixed cells, and propidium iodide, long used as a nuclear marker for flow cytometry which fluoresces in the red range. Far red dyes such as Draq5 can also be used a nuclear counter stain.

Fluorescently labeled phalloidin can be used as a counter stain for actin (Figure 7). This cyclic peptide isolated from the death cap mushroom (Amanita phalloides) has a very high affinity towards F-actin filaments. Because of its relatively small size and chemical stability, fluorescent derivatives are enormously useful in identifying actin filaments with high density labeling of the cytoskeleton which provides a means for the assessment of cellular morphology. As they are commercially available in a multitude of different colors they can be multiplexed with several other cellular probes.

Fluorescently tagged wheat germ agglutinin (WGA) can be used to stain for cell membranes. WGA is a carbohydratebinding protein that selectively recognizes sialic acid and N-acetylglucosaminyl sugar residues which are predominantly found on plasma membrane proteins.

Unlike antibodies, counter stains usually do not have cross reactivity with one another and often can be combined into a single process step - the only limitation would be buffer compatibility. For example, DAPI (5-10 &mug/mL) and phalloidin (20 nM) counter stains can be combined in a mixture of PBS 0.1% Triton X-100 from concentrated stock solution, freshly made on the day of the experiment. A 10-minute incubation at room temperature is sufficient to adequately stain fixed tissue culture cells.

Figure 7. Fixed NIH 3T3 cells expressing GFP counterstained with DAPI and Texas Red labeled phalloidin. NIH3T3 cells cultured in 96-well plates were formaldehyde fixed then counter stained with DAPI and Texas red labeled phalloidin. Cells were imaged with a Cytation&trade 3 using configured with DAPI, GFP and Texas Red light cubes.

Fixation and staining of slides or microplates is a multistep process with a number of reagent additions, wash steps and incubations (Figure 8). The processing of slides or coverslips is generally a low throughput operation and can usually be accomplished with a manual operation. Both fixtures can be treated using small dishes or in the case of slides dedicated Coplin jars can be used to store reagents or as vessels for wash media. Microplates, particularly 96- and 384- well plates with cells are much more amenable to automation. Because microplates have standardized well size, spacing and footprint, automation can be used to provide reagent addition and washing (aspiration and dispense). The small size of coverslips also allows them to be placed in the wells of low-density plates (6- and 12-well) for processing prior to adhering them to a slide.

Figure 8. Automated Workflow for Cell Seeding, Fixation, Permeabilization and Three Color Staining Process. An EL406&trade Combination Washer Dispenser (BioTek Instruments) was used to carry out the process steps for cell fixation, permeabilization and staining with three colors (DAPI, Alexa-Fluor® 488 phalloidin stain, and Texas red labeled secondary antibody) are indicated in red.

Storage of fixed and stained specimens can vary depending on the sample. Specimens on slides (or coverslips) can be treated with a mounting medium and sealed for long term storage. Mounting medium helps preserve the sample and raises the refractive index, which will improve performance with oil objectives. Mounting medium often has added agents to scavenge free radicals and reduce photobleaching. Prolong Gold (Life Technologies) and Fluoromount&ndashG (SouthernBiotech) are examples of commercially available mounting media. It is important to rinse the slide/coverslip in distilled water prior to sealing as the salt from the PBS wash will precipitate out as crystals as it dries. After adhering the coverslip to the slide, allow the slide to dry overnight prior to sealing. Seal the edges of each coverslip with regular transparent nail polish and allow drying for 3 minutes cells are now ready for imaging. Slides should be stored in a light tight container to prevent photobleaching. This will provide semi-permanent preparations that will last months if kept in the dark.

Microplates storage is more challenging. The small well size and well depth makes it almost impossible to seal the sample through the use of a coverslip. Microplates are better stored wet at 4 °C - PBS with 0.1% sodium azide can be used as a preservative. Plates are sealed with an adhesive plate seal and the plates wrapped in aluminum foil. While they will lose resolution over time, clear images can be obtained after one month in storage.

Final Comments

There is no &ldquobest&rdquo procedure for immunostaining and counter staining fixed cells on slides or microplates. There are numerous methods to fix, permeabilize, and stain cells, each with strengths and weaknesses. Immunostaining intracellularly involves detecting small molecules on interior cell structures often using much larger molecules, such as antibodies. Structures need to be stabilized, while at the same time holes need to be opened in membranes large enough to allow antibodies access to the interior. Different antibodies have different affinities and will often recognize different epitopes on the same protein, while different fixative methods will expose or mask different epitopes within the cell. While there is no one-size-fit-all technique for immunostaining, there are a number of different techniques that can be tested in order to optimize results.


Watch the video: Mounting Cells for Fluorescence Microscopy (December 2022).