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Where do lost membrane proteins go after exocytosis?

Where do lost membrane proteins go after exocytosis?


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Exocytotic vesicles take away membrane proteins and glycocalyx on the cell's plasma membrane surface. When those vesicles are released into the interstitial fluid and wherever else, where do they go?

Do they stick to the vesicle everywhere? Or do they get removed from the vesicle? If the vesicle fuses into another cell, do those proteins and carbohydrate attachments stick to the new cell?


In the process of exocytosis materials which are about to be released are transported in small vesicles to the plasma membrane. The plasma membrane fuses with these vesicles and this sets the substances free on the outside of the cell. See the figure (from here):

The other possibility for transport vesicles is that they arrive at their target cell and either fuse with the membrane and release the transported substance into the cell or that they are taken up in another vesicle which is then targeted towards the endosome/lysosome. See the figure (from here):

It doesn't matter which way is chosen, the membrane of the vesicle is recycled either by integration into another membrane of by going through the lysosome.


Cell Biology 04: The Secretory Pathway

The secretory pathway refers to the endoplasmic reticulum, Golgi apparatus and the vesicles that travel in between them as well as the cell membrane and lysosomes. It’s named ‘secretory’ for being the pathway by which the cell secretes proteins into the extracellular environment. But as usual, etymology only tells a fraction of the story. This pathway also processes proteins that will be membrane-bound (whether in the cellular membrane or in the ER or Golgi membranes themselves), as well as lysosomal enzymes, and also any proteins that will live their lives in the secretory pathway itself. It also does some things other than process proteins.

The cytosol and the ‘lumen’ (the liquid that fills the secretory pathway) are different chemical environments, and they normally never mix. The cytosol is reductive (when you’re in the cytosol, you keep meeting molecules that want to offer you electrons), and the ER, Golgi and extracellular environment are oxidative (molecules keep coming up to you asking for electrons). See redox if still confused. This makes for different protein-folding conditions: for instance, disulfide bonds usually only form in oxidative conditions. Moreover, different proteins may live only in the secretory pathway or only in the cytosol. The secretory pathway provides a route for the cell to handle things that might not be good to have in the cytoplasm, and/or are most useful when kept concentrated in a specialized compartment with their desired interacting partners. Hepatocytes (in the liver) sequester drugs and toxins in the smooth ER and break them down for excretion from the body there. The secretory pathway is not contiguous, but every movement between its components is in little bubbled-off microcosms of its own chemical world, called vesicles.

Many proteins that go through the secretory pathway never touch the cytosol – except the parts of membrane proteins that stick out on the cytosolic side. Many of them need chaperones to help with folding, and/or a whole series of post-translational modifications in order to be ready for their native function, and the secretory pathway specializes in providing them all of that.

Today’s lecture will focus on how proteins get translated into the ER and how they travel (in vesicles) between the ER, Golgi and other destinations. This is beautifully depicted in the Life of the Cell video:

The endoplasmic reticulum is the first step in the secretory pathway. Its membrane is continuous with the outer nuclear membrane, though it’s not clear why that matters, since it’s not like proteins begin their life in the nucleus. Rather, mRNAs drift around in the cytoplasm until they get picked up by a ribosome interested in translating them. In ‘posttranslational translocation’ the new protein is moved into the ER after it’s translated. In the more interesting phenomenon called ‘cotranslational translocation’ the ribosome starts translation just like any other protein, but somewhere in the first 16 to 30 amino acids it hits a signal peptide (aka signal sequence). That signal’s motif is often 1 positively charged amino acid followed by 6-12 hydrophobic amino acids. This motif gets recognized by signal recognition particle (SRP, a ‘ribonucleoprotein’ or hybrid RNA/protein molecule) which binds to it and prevents the ribosome from continuing translation. Translation is stopped until the ribosome/SRP complex encounters an SRP receptor on the ER membrane. When they meet, SRP and its receptor each bind one GTP molecule in the ER membrane, which apparently strengthens their interaction. Fortuitously, this all happens adjacent to a Sec61 translocon – a protein complex that forms a channel crossing the ER membrane. The translocon is actually a complex of three different proteins (genes: SEC61A1 or SEC61A2, SEC61B, SEC61G), of which the Sec61a subunit has 10 membrane-spanning a-helices which form the channel. Once the ribosome is docked at the membrane it continues translation, pushing the signal peptide and eventually the whole protein through the channel into the ER lumen. When translation stops, SRP and SRP receptor both hydrolzye their GTP to release each other and the ribosome cargo (this has to require the energy of GTP, since the original binding was downhill), a signal peptidase cleaves the signal peptide off of the nascent protein, and the protein is free to start folding in the ER.

A couple of other players are involved for some ER proteins. Oligosaccharide transferase, which adds glycosyl groups to asparagines in the nascent protein, is part of the translocon complex and it actually performs glycosylation while the new protein is still being translated. So although we call glycosylation a ‘post-translational modification’ it is actually made during translation in this case. Also, to achieve their proper structure, some proteins need to be fully translated before they are allowed to start folding – if the N-terminal portion was allowed to start folding as soon as it entered the lumen, it would end up with the wrong overall structure. To prevent this, sometimes BiP the chaperone binds the protein to keep it unfolded for a while. Imagine BiP as another Pac-Man that bites down on the protein to keep it linear, like Hsc70 in the mitochondrial targeting process (see last week).

The first couple of minutes show the basic scenario described above. Then it moves on to a more complex scenario I’ll introduce in a minute. FYI, the video depicts two ‘controversial’ things not included in the above description: (1) the signal peptide being degraded in the membrane, and (2) a ‘plug protein’ that stops up the channel before/after translation. Not all scientists agree on these two things yet.

All of the proteins that we know go through the secretory pathway were pinpointed there by people doing localization experiments to see where in the cell a protein lies. A weird fact about the ER is that you can put the cell in a blender and afterwards the ER will just start reconnecting to itself, forming little ‘microsomes’ that are not attached to the nucleus but form contiguous bubbles of ER. You can then start to play games with proteases – which break down proteins – and detergents – which solubilize the ER membrane. Assuming your protein of interest is translated, you can check if it (1) survives protease treatment but (2) doesn’t survive protease + detergent treatment, then it’s a secretory pathway protein. The logic is that in case (1) it was protected inside the ER, but in case (2) you dissolved the ER, so it got eaten by the protease. All this assumes you have an antibody or some other way of detecting whether the protein of interest is there after these treatments.

People also used such techniques to figure out that only 70 amino acids of a new protein can be translated before it becomes too late for that protein to end up in the ER. Remember, the signal peptide is in the first 16-30 amino acids, and translocation to the ER depends on SRP being present. Ribosomes translate at a predictable rate, so people got ribosomes started on translating some mRNA and then waited set amounts of time before adding SRP, to see how much translation could occur before SRP could no longer do its job.

The SRP receptor and the Sec61 proteins are ER membrane proteins – and there many other ER membrane, Golgi membrane and lysosome membrane proteins as well. In fact, even the membrane proteins (see class 02) of the cell membrane get processed in the secretory pathway. Many of these have several or tens of transmembrane domains (20-25 hydrophobic amino acids each) that have to be inserted in the correct order and orientation (for example, you really want your ion channels and transporters pointed in the right direction, into vs. out of the cell). Accordingly there are a bunch of fancy biological mechanisms for getting these proteins inserted into the membrane correctly. This is what the latter half of the above video depicts.

So here’s a tautology: some proteins have a topogenic sequence which determines their orientation in the membrane. This sequence is made of two types of signal sequences:

  • a stop-transfer sequence (abbreviated STA for some reason) is a 22-25 hydrophobic amino acid sequence somewhere in the middle of the protein that forms an alpha helix. When encountered it gets shoved into the membrane, and then translation of the rest of the protein continues in the cytosol. So this kind of ‘undoes’ the translocation to the ER that was started by the signal peptide at the beginning (N terminus) of the protein.
  • a signal anchor sequence (abbreviated SA) is also a 22-25aa hydrophobic alpha helix, but with a series of

With those two signals as building blocks, you can imagine a protein with a series of stop transfer and signal anchor sequences to create a whole series of back and forth transmembrane domains stitched into the membrane as if by a sewing machine. People have classified the membrane proteins into five categories:

  1. Type I has just a signal peptide and then one stop transfer in the middle. Therefore it ends up with its (hydrophilic) N terminus in the lumen, its (hydrophobic) middle in the membrane and its (hydrophilic) C terminus in the cytosol.
  2. Type II does not start with a signal peptide. It starts out like any other protein, but in the middle it has a signal anchor sequence with the +++ amino acids coming first and the hydrophobic series after. This makes the protein get translocated midway through translation, with the already-translated N-terminal part sticking out into the cytosol (since the +++ have to stay cytosolic) and the now-beginning-to-be-translated C-terminal part getting translated directly into the ER. So it ends up transmembrane with its C terminus in the ER and N terminus in the cytosol – opposite of Type I.
  3. Type III is like Type II – no signal peptide, just a signal anchor in the middle, but in this case the +++ come after the hydrophobic sequence, which reverses the orientation. So this ends up with its N terminus in the ER and its C terminus in the cytosol. Opposite of Type II and, in the end, the same as Type I, though it got there in a different way – it does not have a signal peptide that gets cleaved off in the ER.
  4. Type IV or ’multipass’ proteins have an alternating series of signal sequences and stop transfer sequences. These are clearly more than one ‘type’, yet are not nearly as diverse as your combinatoric imagination might allow. The orientation of the first signal sequence determines whether the N terminus will end up in the cytosol or ER, and total number of stop transfer + signal anchor sequences determines where the C terminus will end up: an even number = same side as N terminus, odd number = opposite side as N terminus. The STA and SA sequences have to strictly alternate, with the exception that you can start with two signal anchor sequences if the first one is oriented with the N terminus into the cytosol. Just to make a mockery of this categorization scheme, people have defined some incompletely-defined subtypes of Type IV, where Type IVa is N-terminal in cytosol (thus it starts like a Type II protein) and Type IVb is N-terminal in the lumen (it starts like a Type III protein but then has another SA sequence that puts it back into the ER). GLUT1 from Class 02 is a Type IVa. -anchored proteins, which are the fifth type but aren’t called Type V, start with a signal peptide and end with a hydrophobic C-terminus which stays embedded in the membrane. That hydrophobic end gets cleaved off and replaced with GPI, which also stays embedded in the membrane. PrP is one of these – more on that later.

By now we’ve discussed how proteins can end up in the ER lumen or spanning the ER membrane. Most proteins leave the ER within minutes, transported in vesicles bound for the Golgi and then later for excretion, lysosomes or the cell membrane. That forward direction of travel is called anterograde going backwards from Golgi to ER is retrograde transport.

Both types of transport take place in membrane-bound vesicles. These bud off of the membrane of wherever they’re coming from, and later fuse to the membrane of wherever they’re headed – beautifully depicted at

2:25 in the Life of the Cell video above. The body from which the vesicles form is the ‘donor compartment’, and the destination they later fuse to is the ‘acceptor compartment’.

The budding process requires that G proteins in the membrane recruit Coat proteins. Specifically, for anterograde transport, G protein Sar1 (gene: SAR1A) recruits COPII (‘cop two’) for retrograde transport, an ARF G protein recruits COPI (pronounced ‘cop one’). These G proteins are activated to do this job when GEF loads them with GTP, swapping out GDP.

So the steps in anterograde transport, for example, are as follows:

  1. Sec12-GEF (Sec stands for secretory) loads Sar1 with GTP. When bound to GDP, Sar1 just floats around the donor compartment, but when bound to GTP, it undergoes conformational change that causes its otherwise-buried N-terminal hydrophobic tail to protrude, making it stick into the membrane, where COPII proteins then start to accumulate because they really like that tail.
  2. The COPIIs start to polymerize and, due to its conformation, have an intrinsic preference for curvature, so their accumulation starts to make budding happen. At the same time, membrane bound proteins that need to be transported – identified by a DXE (i.e. aspartate-anything-glutamate) amino acid sequence that forms a binding site in their cytosolic part – get recruited to the newly forming vesicle. Membrane-bound proteins act as receptors, recruiting lumenal proteins that are bound for the Golgi to hang out in the concave space where they’ll end up in the vesicle once it forms.
  3. Once enough COPII have arrived, the vesicle buds off, at which point Sar1 hydrolyzes its GTP, providing the energy for it to suck its hydrophobic tail back into itself, cutting the COPIIs loose. The vesicle is now disconnected from the donor compartment.
  4. Now, for poorly explained (or poorly understood?) reasons, the coat of COPIIs just disassembles, exposing receptors under the coat which direct the targeting of the vesicle. Once the vesicle arrives at its destination, Rab-GTP embedded in the vesicle membrane interacts with a Rab effector embedded in the acceptor compartment membrane. A sideways glance is exchanged, interest is kindled. Soon the vesicle will fuse to the membrane. proteins present on both the v esicle and t arget membrane (V-SNARE and T-SNARE respectively) interact to bring the membranes even closer. In this example we’ll consider VAMP (the VAMP_ genes) as the V-SNARE and Syntaxin (the STX__ genes) and SNAP25 (SNAP25 gene) as the T-SNAREs. Syntaxin and SNAP25 are both membrane proteins Syntaxin has 1 alpha helix and SNAP25 has 2, all on the cytosolic side. The alpha helices drive the interaction with VAMP. The opposing sides’ alpha helices have extremely strong affinity for one another, bringing the membranes close enough to fuse. Once this has happened, prying the V-SNAREs and T-SNAREs apart again requires two proteins: NSF (gene: NSF stands for NEM sensitive factor) and alpha-SNAP (gene: NAPA), a soluble NSF attachment protein. NSF is an ATPase, and burns ATP to drive the energetically uphill disassembly of the complex.

Now for retrograde transport. Why is there retrograde transport at all? Here is a non-exhaustive list of some reasons:

  • Some membrane proteins start their life in the ER, need to get modified in the Golgi, but then need to get back to the ER. They do this with a KKXX amino acid sequence.
  • There’s also a KDEL amino acid sequence at the C terminus of some lumenal proteins which is suppsoed to keep them in the ER, but it’s not perfect – sometimes they end up in the Golgi, in which case they’re targeted back to the ER via retrograde transport dependent on that KDEL sequence for recognition. The mechanism is kind of neat – the proteins that recognize and bind to KDEL do so only at low pH, and the pH of the Golgi is lower than the ER, so they bind KDEL in the Golgi, then release it when they’re back in the more neutral pH of the ER.
  • Also, think about it, all the proteins that participate in anterograde transport – the V-SNARES, Rab, etc. – have to get back to the ER so they can do it all over again, like how the bus has to get back to the bus depot at the end of the day.
  • As we’ll see shortly, the Golgi come in multiple stages which depend on the addition of enzymes from further downstream.

The process of retrograde transport is not so different from anterograde. It uses ARF instead of Sar1, COPI instead of COPII, but it works the same: ARF loaded with GTP lets its hydrophobic tail stick into the membrane, attracting the attention of COPIs. COPI has two components, COPIalpha and COPIbeta, both of which interact with that KKXXX sequence to recruit membrane-bound proteins destined for retrograde transport. Some proteins also have an RR sequence (anywhere in the protein) which can flag them for retrograde transport.

The Golgi apparatus is not contiguous. It is a stacked set of separate subcompartments called sacs or cisternae. Different compartments have different properties and proteins visit them in a particular order. In order from ER to cell membrane, the Golgi compartments are called cis, medial, trans and trans-Golgi network. Each compartment has different enzymes that modify proteins, and the modifications have to happen in a certain order, hence the need for a stacked set of compartments.

But as proteins mature in the Golgi, it’s not as though they bud off in vesicles from one compartment and move to the next. Rather, the compartment they are already in moves outward and ‘matures’ as new enzymes are added to it (from further down the Golgi chain) via retrograde transport. Weird, right? It’s kind of like if instead of moving from an elementary school to a middle school to a high school you just stayed in one school building for your whole childhood and adolescence, and they just brought in new textbooks and teachers every year to keep it appropriate to the grade that you and your classmates had now reached. Here’s what the Golgi look like as they move and evolve:

So there’s (little or) no anterograde transport within the Golgi, but plenty of retrograde transport to bring each new round of enzymes in. When proteins have finally completed the full K-12 curriculum of the Golgi network, they do undergo transport to move on to their final destinaton. They bud off in a vesicle which will go one of three places:

    – fusion with the cell membrane. Thus the lumenal proteins will be secreted extracellularly, and the membrane proteins will become cell membrane proteins. – these just stick around as vesicles in the cell until needed – where ‘needed’ means they do eventually undergo exocytosis. In neurons, this is where neurotransmitters are stored until an action potential demands their secretion into the synapse. In the stomach, the cells that produce gastric enzymes keep those enzymes in secretory vesicles until food intake triggers their release into the stomach. - where misfolded proteins go to get degraded.

The transport from the trans-Golgi network on to these destinations is different from the other transport discussed above and often involves clathrin (CLT__ genes). Vesicles budding off have a two-layer coat, with a dapter p rotein (AP) complexes as the inner layer and clathrin as the outer layer. The adapter proteins have a target signal with a YXXh motif (h = Φ = any hydrophobic amino acid). Clathrin forms the so-called ‘clathrin-triskelion’ formation shown here:


(Image thanks to Wikimedia Commons user Phoebus87)

Clathrin is also responsible for endocytosis – budding off of vesicles of extracellular stuff (and cell membrane proteins) to come into the cell. This is called clathrin-mediated endocytosis. Receptors in the cell membrane get endocytosed very frequently: the whole population of hormone receptors turns over about every hour, especially when hormones are being received. Taking up the receptor into a vesicle is one way for the cell to cut off the incoming signal until it can be processed.
The plasma membrane notes discuss cystic fibrosis briefly: CFTR is an ABC transporter responsible for pumping Cl - out of the cell (it also lets Na + in). Loss-of-function mutants don’t pump Cl - , which removes the driving force for osmosis, thickening the mucus and causing breathing problems. There are at least 127 different loss-of-function CFTR mutants (at least, that’s how many Natera tests for) that (if both alleles are disabled) cause cystic fibrosis. The most common mutation is ΔF508, which is

3% of all European CFTR alleles and about 70% of mutant ones. The loss of that one phenylalanine changes CFTR’s conformation so that the di-acidic exit code (amino acids D565 and D567) that targets CFTR for exocytotic vesicles is no longer correctly exposed and the protein never makes it to the cell membrane [Wang 2004].

discussion section

In section we read Hu 2009, who showed that atlastin proteins are involved in creating the tubular ER network. The evidence came almost entirely from protein-protein interactions. I was surprised this paper was a big deal, because there have been a million papers showing protein-protein interactions for huntingtin, and no one really believes all of them and it hasn’t necessarily gotten us any closer to knowing what huntingtin does or what goes wrong in Huntington’s Disease. But apparently Hu was able to make a pretty clean case for the atlastins’ interactions with reticulons as implying a role in ER formation. It helps that Hu was able to show a ‘genetic interaction’ in addition to a physical (binding) interaction. A ‘genetic interaction’ (I had to look it up) means when “Sometimes mutations in two genes produce a phenotype that is surprising in light of each mutation’s individual effects. This phenomenon, which defines genetic interaction, can reveal functional relationships between genes and pathways.” [Mani 2007].

This is a decade old, so some stuff may be outdated, but I found Harris 2003 (ft)’s review of PrP cell biology extremely clear and helpful. Kim & Hegde 2002 was also helpful. PrP is a secretory pathway protein. Its first 22 amino acids (MANLGCWMLVLFVATWSDLGLC) are a signal peptide that causes cotranslational translocation to the ER. Normally, PrP just gets GPI-linked at its C terminus and is anchored to the exoplasmic side of the membrane. But amino acids 111-134 (HMAGAAAAGAVVGGLGGYMLGSAM) are a sort of weak signal anchor sequence (Type II, with the +++ amino acids coming before the signal anchor) that sometimes but not always becomes a transmembrane domain, inverting the C terminus into the lumen. Even more confusingly, that sequence can sometimes just end up as a transmembrane domain without the inversion, so that the N terminus is in the lumen. So there are three membrane topologies of PrP: regular old GPI-anchored, and two transmembrane orientations, as depicted in Harris 2003 Fig 3:

Note how weird Ctm PrP is. It’s transmembrane yet also GPI-anchored, and the N-terminal signal peptide is never cleaved off. Normally, the transmembrane forms are < 10% of total PrP. In some laboratory conditions the percentage is higher, and two of the GSS-causing mutations (A117V and P105L) also increase the fraction of Ctm PrP to 20-30% of all PrP. Of these three forms, there is a good amount of evidence that Ctm PrP is toxic, and that it might play a role in prion formation, though most genetic prion disease mutations (including FFI D178N) do not appear to affect the membrane topology of PrP or the fraction of Ctm PrP.

After PrP goes through the Golgi, it is targeted for the cell membrane. But according to Harris, it doesn’t just sit there – it frequently through clathrin-mediated endocytosis and cycles through the cell every

60 minutes, with some molecules being cleaved on each cycle. Copper stimulates this endocytosis of PrP. Most genetic prion disease mutations change the localization of PrP – usually when a mutation is present, less PrP is found on the cell surface, with more accumulating in the ER.

About Eric Vallabh Minikel

Eric Vallabh Minikel is on a lifelong quest to prevent prion disease. He is a scientist based at the Broad Institute of MIT and Harvard.


REVIEW article

  • Key Laboratory of Algal Biology, Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan, China

Cilia and flagella are highly conserved organelles in eukaryotic cells that drive cell movement and act as cell antennae that receive and transmit signals. In addition to receiving and transducing external signals that activate signal cascades, cilia also secrete ciliary ectosomes that send signals to recipient cells, and thereby mediate cell�ll communication. Abnormal ciliary function leads to various ciliopathies, and the precise transport and localization of ciliary membrane proteins are essential for cilium function. This review summarizes current knowledge about the transport processes of ciliary membrane proteins after their synthesis at the endoplasmic reticulum: modification and sorting in the Golgi apparatus, transport through vesicles to the ciliary base, entrance into cilia through the diffusion barrier, and turnover by ectosome secretion. The molecular mechanisms and regulation involved in each step are also discussed. Transport of ciliary membrane proteins is a complex, precise cellular process coordinated among multiple organelles. By systematically analyzing the existing research, we identify topics that should be further investigated to promote progress in this field of research.


Results

Imaging single synaptic microvesicles

Here we imaged single synaptic-like microvesicles in living cells with total internal reflection fluorescence (TIRF) microscopy 5 . Specifically, we used a microvesicle-targeted pH-sensitive fluorescence probe (VAChT-pH) based on the VAChT (Fig. 1a) 6 . Single vesicles containing this probe brighten when the fusion pore of the vesicle opens after exocytosis and the acidic lumen of the vesicle is neutralized by the extracellular buffer 6 . Figure 1b shows two cells expressing VAChT-pH. Fluorescence was scattered across the bottom surface of the cell, where it was confined to small puncta. To test if these puncta were on the outside of the cell, we superfused cells with a low pH solution (pH 5.5 Supplementary Fig. S1). A dramatic dimming of the cells was measured during this treatment (Supplementary Fig. S1a-c). Single VAChT-pH puncta dimed and then re-brighted, indicating that many of the puncta were on the extracellular face of the plasma membrane. Some puncta did not dim, indicating that they were in intracellular compartments. To test if VAChT-pH was contained in acidic compartments, we superfused cells with ammonium chloride (Supplementary Fig. S1d-f). This chemical breaks down intracellular pH gradients. Cells and some fluorescent puncta exposed to this solution brightened, indicating that some VAChT-pH is located in intracellular acidic compartments (Supplementary Fig. S1d-f). Combined, these results indicate that VAChT-pH was present both in clusters on the plasma membrane and in acidic compartments within the cell.

(a) Cartoon of the microvesicle probe VAChT-pH. (b) Image of two PC12 cells expressing VAChT-pH imaged with TIRF. Scale bar equals 5 μm. (c) Frames from a movie where a single VAChT-pH-containing vesicle undergoes depolarization-triggered exocytosis, and (d) the corresponding fluorescence from the centre 750 nm-radius circle of that region. Scale bar equals 2 μm. (e) The mean VAChT-pH fluorescence from triggered exocytic vesicles (83 events, 13 cells). (f) Cartoon of the ratiometric pH probe VAChT-pH-mCherry. (g) The mean VAChT-pH-mCherry fluorescence from triggered exocytic vesicles in both the pHluorin and mCherry channels (36 events, 3 cells). The ratio of these two intensities is shown in h. Error bars are s.e.m. Norm. fluor., normalized fluorescence.

To evoke exocytosis, we depolarized cells with high potassium. This solution induced rapid and numerous exocytic events. Bright flashes could be seen across the bottom surface of the cell. These events were rare in unstimulated cells. Figure 1c shows an example event (Supplementary Movie 1). Ten seconds before exocytosis, the vesicle is not visible, but when the fusion pore opens, a bright flash occurs within one frame (500 ms) and creates a bloom of fluorescence that radiates outward in all direction and dims (Fig. 1c). After exocytosis, some of this fluorescence was captured on preformed VAChT-pH clusters near the exocytic site (Fig. 1c). The fluorescence from the local membrane area (750 radius circle) surrounding this vesicle is plotted in Fig. 1d. This trace shows that VAChT-pH increases at exocytosis, decays within a few seconds and then plateaus to an intensity close to half of the peak value. The corresponding fluorescence from many events (83 events, 13 cells) is plotted in Fig. 1e. On average, VAChT-pH brightens rapidly and then decays. There is, however, a substantial amount of fluorescence (61±0.03%) that remains near the site of fusion (Fig. 1e).

To determine if the VAChT-pH decay was due to loss of the protein into the plasma membrane (full-fusion) or was due to the direct recapture and re-acidification of the vesicle (kiss-and-run), we tagged VAChT-pH with mCherry. VAChT-pH-mCherry, positions the pH-sensitive pHluorin inside the vesicle lumen and the pH-insensitive mCherry into the cytoplasm (Fig. 1f). This sensor acts as a ratio-metric probe for pH changes in vesicles. The green fluorescence from VAChT-pH-mCherry rapidly brightens and then decays to 58±0.07% (Fig. 1g). Similarly, the mCherry fluorescence increases and then decays at the same rate, and to the same extent as pHluorin (62±0.07% Fig. 1g). The ratio of the red-to-green is shown in Fig. 1h, and shows that after exocytosis there is no change. Thus, the decay in VAChT-pH after exocytosis is not due to direct recapture and acidification (kiss-and-run), but instead is due to the loss of the probe into the membrane. These results support a full-fusion mechanism of exocytosis for VAChT.

To analyse the spread of VAChT-pH, we measured four radial line scans centred over vesicles (Fig. 2a). These scans generated an average one-dimensional kymograph of the two-dimensional membrane surrounding exocytic events. Figure 2a shows results of averaging the kymographs of many VAChT-pH fusion events together. Upon exocytosis, fluorescence brightens and spreads rapidly. Fluorescence, however, progressed only a short distance. As seen in the example in Fig. 1c, some of the diffusing fluorescence remains near the site of fusion (Fig. 2a). The fluorescence from these kymographs is plotted in Fig. 2b. Analysis of these data is shown in Fig. 2c, where two-dimensional Gaussian functions were fit to vesicles. This plot shows that VAChT initially spreads rapidly, but remains relatively constant in size 10 s following exocytosis. We conclude that microvesicles release VAChT into the plasma membrane. Most of this material, however, does not spread more than a few hundred nanometres (σ-values at 30 s, 1.1±0.1 μm) from the site of exocytosis. These results are consistent with several models of release for VAChT, which we consider below.

(a) Mean normalized radial linescans of microvesicles during exocytosis (46 events, 13 cells). Scale bar equals 1.5 μm. (b) Intensity plot of the same radial linescans. (c) Mean σ-value of a Gaussian function fit to each of the events used for a and b. Error bars are s.e.m.

Vesicle material is trapped in clathrin-coated structures

The cytosolic tail of VAChT binds to the adaptor protein AP2 (refs 7, 8). To determine if endocytic proteins were responsible for clustering vesicle material on the cell surface, and are involved in the retention of VAChT near sites of exocytosis, we imaged VAChT-pH together with two endocytic markers. Figure 3a shows an image of VAChT-pH and a marker for clathrin-coated structures (clathrin light chain tagged with dsRED, mLC-dsRED) 9 . These images show that the bottom surface of PC12 cells have a large number of clathrin-coated structures (0.95 clathrin spots per μm 2 ±0.02 spots per μm 2 , n=11 cells, 79 regions (72 μm 2 area)). A similar density for clathrin was found with antibody staining (Supplementary Fig. S2). To test if MLC-dsRED labelled all the clathrin structures, we immunostained cells expressing MLC-dsRED with antibodies against the heavy chain of clathrin. Almost all the antibody-stained clathrin spots were labelled with MLC-dsRED (Supplementary Fig. S3). Furthermore, the number of antibody-stained clathrin structures was not increased compared with untransfected controls (X22 density of MLC-dsRED-transfected cells: 1.05±0.02 clathrin spots per μm 2 , X22 density of control cells: 1.06 clathrin spots per μm 2 ±0.03 Supplementary Figs S2 and S3). These results show that the density of clathrin was not changed by the expression of clathrin light chain (Supplementary Figs S2 and S3).

(a) TIRF image of a cell co-expressing VAChT-pH and MLC-dsRED along with the overlay image. Scale bar equals 5 μm. (b) Mean extracted and normalized square regions centred on VAChT-pH spots and the corresponding regions from the MLC-dsRED channel (n=11 cells, 1,374 regions). Scale bar equals 2 μm. (c) The two-dimensional correlation values between the images used in b. (d) TIRF image of a cell co-expressing VAChT-pH and AP2-mCherry along with the overlay image. Scale bar equals 5 μm. (e) Mean extracted and normalized square regions centred on VAChT-pH spots and the corresponding region from the AP2-mCherry channel (n=6 cells, 964 regions). Scale bar equals 2 μm. (f) The two-dimensional correlation values between the images used in e. Avg, average.

When green images (Fig. 3a) were compared with red, a large fraction of VAChT-pH and clathrin colocalized (Fig. 3a). To analyse colocalization, we extracted 1,374 small (8.5 × 8.5 μm) images from 11 cells. In these regions, the centre of VAChT-pH structures were aligned to the middle pixel of the image. The corresponding regions from the MLC-dsRED image were also extracted. The fluorescence from these regions were normalized and averaged. This analysis showed a small distinct spot in the VAChT-pH image that co-localized with clathrin (Fig. 3b). The correlation between these regions is plotted in Fig. 3c and shows a strong and positive correlation. Visual inspection of 215 randomly chosen regions indicated that 85% of the central VAChT-pH spots had a corresponding signal in the MLC-dsRED channel. Similar analysis was done with VAChT-pH and AP2-mCherry (Fig. 3d-f). These images and their analysis show that, similar to clathrin VAChT-pH and AP2-mCherry strongly colocalize. Average VAChT-pH and AP2-mCherry image pairs have similar correlation values, sizes and distributions (Fig. 3d-f). Again, visual inspection of 215 randomly chosen regions showed that 82% of all the central VAChT-pH spots colocalized with AP2-mCherry. To test for the association of clathrin and AP2, we measured the colocalization of MLC-dsRED and AP2-green fluorescent protein (GFP Supplementary Fig. S4). Visual inspection indicated that 98% of the MLC-dsRED had a corresponding signal in the AP2-GFP channel. Thus, almost all of the MLC-dsRED structures contained AP2-GFP. We conclude that there is a resting pool of VAChT-pH on the surface of the cell that is clustered on endocytic structures composed of clathrin and AP2.

After exocytosis, vesicle material is trapped over clathrin

We next asked if the corralling of VAChT-pH observed after exocytosis is related to clathrin. To answer this question, we triggered exocytosis in cells expressing VAChT-pH and MLC-dsRED. We then imaged the release of VAChT-pH from single exocytic vesicles and compared these images with clathrin. Figure 4a shows a single exocytic vesicle arriving at the surface and rapidly brightening. VAChT-pH left the site of exocytosis and diffused radially into the plasma membrane where a fraction of the fluorescence was rapidly (<1 s) captured on a preformed clathrin structure (Fig. 4a Supplementary Fig. S5 and Supplementary Movie 2). This trapped VAChT-pH remained associated with clathrin for over 20 s.

(a) Sequential two-colour TIRF images of a single vesicle fusing with the plasma membrane. The location of fusion is indicated by the blue arrow. After fusion, VAChT-pH is captured on a preformed clathrin-coated pit (yellow arrow) located close to the site of exocytosis. Scale bar equals 2 μm. (b) Mean radial linescans through images centred on clathrin structures located within 1 μm of an exocytic event (n=7 cells, 35 regions). Fusion occurs at zero seconds. Scale bar equals 2 μm. (c) The measured fluorescence from VAChT-pH and MLC-dsRED from a circle with a radius of 375 nm centred on the clathrin structures shown in b. (d) Mean radial linescans through images centred on clathrin-free areas located within 1 μm of an exocytic event (n=7 cells, 43 regions). (e) The measured fluorescence from VAChT-pH and MLC-dsRED from a circle with a radius of 375 nm centred on the centre of the clathrin-free areas shown in d. Spatial analysis comparing sites of exocytosis (f, yellow cross) to clathrin (f, red circles). Scale bar equals 2 μm. The blue circle indicates an area 1 μm from the centre of exocytosis. (g) Plot of the number of clathrin structures located within 1 μm of fusion events (n=11 cells, 79 regions, 4,383 clathrin structures). (h) Histogram showing nearest-neighbour distances between fusion sites and clathrin spots. (i) Histogram showing the nearest-neighbour distances between random sites and clathrin spots. Error bars are s.e.m. Avg. fluor., average fluorescence Norm. fluor., normalized fluorescence.

To analyse the capture of VAChT-pH on clathrin, we measured fluorescence around clathrin structures located within1 μm of fusion events. The average kymographs centred over clathrin is shown in Fig. 4b. From this analysis, it is evident that clathrin is stable at the membrane before exocytosis. At the moment of fusion, VAChT-pH brightens in the membrane area surrounding clathrin, steadily increases and is then focused onto the central clathrin-containing spot (Fig. 4b). We observe that the width of the VAChT-pH spot decrease after exocytosis and aligns with the central spot of clathrin in the kymograph (Fig. 4b). A plot of the fluorescence intensity during exocytosis measured in a small membrane area (375 nm radius circle) over clathrin is shown in Fig. 4c. In these clathrin-containing regions, VAChT-pH fluorescence steadily increases after exocytosis and then plateaus (Fig. 4c). The small and slow decline of fluorescence over clathrin during the plateau phase could be due to several experimental issues, including photobleaching, vesicle movement, re-acidification and endocytosis. These data demonstrate that the trapping of VAChT-pH occurs over resident clathrin structures located within a micron of exocytosis.

To show that VAChT-pH is specifically trapped over clathrin after exocytosis, we measured VAChT-pH in regions that did not contain clathrin and were located within 1 μm of fusion events. The average kymograph centred over clathrin-free areas is shown in Fig. 4d. In these regions, VAChT-pH brightens at exocytosis and is then rapidly lost. Specifically, VAChT-pH diffuses from the central, clathrin-free area, and is then captured on the surrounding clathrin. Unlike Fig. 4c, the fluorescence from clathrin-free areas (375 nm radius circle) increases at exocytosis, does not continue to increase and then rapidly decays (Fig. 4e). These data demonstrate that after exocytosis, VAChT-pH freely diffuses in membranes that do not contain clathrin, and is caught on membranes that do contain clathrin.

To test for a spatial coupling between sites of exocytosis and endocytosis, we mapped the location of 79 exocytic vesicles in relation to all clathrin spots. Figure 4f shows an example image of a field of clathrin where an exocytic event occurs in the centre (Fig. 4f, yellow cross). For this analysis, we determined the location of all the clathrin structures in each image surrounding exocytic sites (11 cells, 79 regions, 4,383 spots). The same analysis was done for control images from the same cells (7 cells, 67 images, 3,941 spots). This analysis shows that there is a dense network of clathrin across the cell surface. Interestingly, there was no increase in the number of clathrin structures near sites of exocytosis (2.9 spots±0.2 spots) compared with the control images (2.7 spots±0.2 spots Fig. 4g). A histogram showing the distance between exocytic sites and the closest clathrin structure is plotted in Fig. 4h. The closest clathrin structure to each fusion site was 482±27 nm. Figure 4i shows that the nearest-neighbour spacing of clathrin structures is 706±11 nm (4,373 structures, 79 regions, 11 cells). For controls, the clathrin structure nearest to the centre of the image was 435±23 nm and the nearest-neighbour spacing of clathrin was 711±11 nm. Thus, material exiting a vesicle needs to go no further than a few hundred nanometres before it encounters a preformed clathrin structure. This distribution of clathrin, however, is not linked to sites of exocytosis. Instead, it is randomly generated by the spacing and high density of resident clathrin structures.

To support the hypothesis that classical clathrin-mediated endocytosis was linked to the surface distribution of VAChT, we treated cells with the dynamin inhibitor dynasore 10 . Like MLC-dsRED and AP2-mCherry, Dynamin1-mCherry (Dyn1-mCherry) co-localized with VAChT-pH (Supplementary Fig. S6). Visual inspection indicated that 64% of VAChT-pH spots co-localized with Dyn1-mCherry spots (Supplementary Fig. S6). When cells were exposed to 80 μM dynasore for 30 min, a dramatic increase in surface fluorescence over time was observed (Supplementary Fig. S6). This increase in fluorescence occurred in both the stable fluorescent puncta and plasma membrane regions not associated with puncta. Thus, the surface distribution and recycling of VAChT-pH is dependent on the action of components of clathrin-coated pits, namely dynamin.

Architecture of clathrin determined by iPALM

From the above data, we conclude that a large fraction of vesicle material is rapidly and locally captured over pre-formed clathrin structures after triggered exocytosis. Standard optical microscopy (for example, TIRF) is restricted by the diffraction limit of light. This fact limits the resolution of cellular structures to

200 nm with TIRF. Thus, we used a super-resolution optical technique known as interferometric photo-activation localization microscopy (iPALM) to image the three-dimensional sub-diffraction size and distribution of clathrin in PC12 cells 11 . To accomplish this, we tagged clathrin light chains with the photo-switchable fluorescent protein mEOS2 (mLC-mEOS2 Fig. 5a) 12 . iPALM was used to reconstruct the size, shape and distribution of clathrin at the bottom surface of the plasma membrane. iPALM also allows one to analyse the distribution of clathrin molecules in the plane perpendicular to the coverslip 11 . Figure 5b shows a single cell imaged with iPALM. The images to the left shows the cell imaged with a conventional TIRF, and to the right shows the same cell imaged with iPALM. Clearly, individual spots of clathrin are difficult to distinguish with conventional imaging. However, distinct clathrin structures are clearly visible with iPALM.

(a) Cartoon of a single clathrin triskeleon with a light chain labelled with a fluorescent protein. The triskeleons assemble together to form clathrin-coated structures. (b) Conventional TIRF optical image and (c) iPALM measurement of MLC-mEOS2 in a PC12 cell. Scale bar equals 5 μm. In b, the blue arrows point to gold particles used for alignment. In both cases, a magnified view of the white box is shown below. Scale bar equals 1 μm. (d) Histogram of clathrin spacing in PC12 cells measured with iPALM. (e) Histogram showing the width of the same structures. (f) Plot of width versus depth of the clathrin objects measured with iPALM. Error bars are s.e.m.

From images of entire cells, we were able to calculate the sub-diffraction distribution of clathrin. The total density of resolvable spots was 2.00±0.09 spots per μm 2 (3 cells). Nearest-neighbour analysis showed that the average spacing was 560±1 nm (n=3 cells, 305 spots Fig. 5d). This was 200 nm closer than we observed with conventional TIRF. To systematically analyse the shape of all the structures on the surface, we measured the width and depth of each structure from three cells in the x, y and z dimensions relative to the glass coverslip. Figure 5e shows that the size of clathrin-coated structures ranged from 50 to 450 nm in diameter (mean: 224±5 nm, n=3 cells, 401 structures). These structures varied in shape with 42% less than 75 nm in depth (Fig. 5f). From these measurements, there was no strong correlation between the width and depth of individual pits. These data support the finding that the density of clathrin structures in these cells is very high. Furthermore, individual structures were either flat or domed. The functional importance of these two types of structures is yet to be determined. Finally, the two-fold increase in density observed with iPALM can be explained by the inability of traditional optical imaging (TIRF) to delineate between two closely spaced structures and one large structure.

Architecture of clathrin determined by electron microscopy

To further map the architecture of clathrin, we performed electron microscopy. We first prepared platinum-shadowed membrane sheets from PC12 cells 13 . We then used whole-cell transmission electron microscopy to visualize the entire inner surface of the plasma membrane (Supplementary Fig. S7). We observed two types of clathrin structures: (1) distinctive flat patches and (2) domes, or pit-like structures (Supplementary Movie 3). Figure 6a shows four regions where both flat patches and pits of clathrin can be seen. To determine the shape of these structures, we measured the minimum and maximum dimension (1,045 clathrin structures, 96 regions, 3 cells). The structures were further classified by eye as flat or domed. Figure 6b shows a scatter plot of these measurements. The structures have a maximum diameter of 129±1 nm (1,045 clathrin structures, 96 regions, 3 cells). Figure 6c show this nearest-neighbour analysis. Clathrin structures were distributed at a nearest-neighbour density of 463±8 nm (3 cells). The total density was 1.6 clathrin structures per μm 2 ±0.11 (96 regions, 3 cells). To further confirm these densities, we performed super-resolution imaging (Supplementary Fig. S8). The density observed with ground-state depletion (GSD) imaging showed 2.3 structures per μm 2 ±0.08 (92 regions, 6 cells). Again, these data support the finding that the density of clathrin on the surface is very high, between 2 and 11 times greater than previously reported from other cell types 14,15,16,17,18,19,20 . Furthermore, these data demonstrate that the density of clathrin structures measured in iPALM was not a consequence of overexpression. The differences between our EM, iPALM and GSD-measured clathrin densities likely resulted from the systematic measurement errors of each method. For example, some clathrin structures are hidden by other cellular structures in our EM images. Furthermore, background fluorescence could result in a slight overcounting of clathrin structures in iPALM and GSD imaging. These measurements support the model in which fusion occurs randomly among sites of endocytosis, and diffusing material can be rapidly captured on preformed clathrin-coated structures waiting on the cell surface to trap material destined for endocytosis.

(a) Four representative transmission electron microscopic images of the inner surface of the plasma membrane of PC12 cells. A representative flat (yellow arrow) and domed (pink arrow) structure are indicated. Scale bar equals 200 nm. (b) Plot of minimum and maximum caliper measurement for all visible clathrin structures over the entire cell (n=3 cells, 64 regions, 1,045 clathrin structures). Pink circles are structures classified as domed and purple circles are structures classified as flat. (c) Nearest-neighbour analysis of all the clathrin structures shown in b.

Modelling

To test the hypothesis that a dense field of traps can limit the spread of VAChT, we developed a simple physical model (Fig. 7a). First, particles in the centre of a small field were allowed to undergo random two-dimensional Brownian diffusion. These simulations were observed to lose all the particles rapidly (Fig. 7b and Supplementary Movie 4). When we introduced a population of traps into the simulation (1, 2 and 5 traps per μm 2 ), we observe a rapid capture of particles near sites of release (Fig. 7b and Supplementary Movie 5). The amount of capture was directly related to the density of traps. This is observed in Fig. 7b and Supplementary Movie 5. A plot measuring the fluorescence from simulations shows that without traps fluorescence is rapidly lost from the centre (750 nm radius region) into the surrounding membrane (Fig. 7c). The plot, however, of average simulations that included traps shows a rapid loss and then a plateau of fluorescence (Fig. 7c). The size of the plateau is proportional to the number of traps. These simulations support the hypothesis that a dense network of traps can account for the limited spread and rapid capture of VAChT. Furthermore, the density of traps can modulate the amount of material lost or remaining near any individual site of release.

(a) Cartoon of the model for the release and capture of VAChT. (b) Radial kymograph of fluorescence from the mean of five diffusion simulations. The simulations contained 0, 1, 2 and 5 traps per μm 2 . The material rapidly exits the centre, but a large amount of the particles are caught and remain in the centre in simulations that contain traps. Scale bar equals 2 μm. (c) Plot of the mean intensity contained in circular regions with a radius of 750 nm from the above simulations. Error bars are s.e.m.


Some of the integral membrane proteins that a cell displays at its surface are receptors for particular components of the ECF. For example, iron is transported in the blood complexed to a protein called transferrin. Cells have receptors for transferrin on their surface. When these receptors encounter a molecule of transferrin, they bind tightly to it. The complex of transferrin and its receptor is then engulfed by endocytosis. Ultimately, the iron is released into the cytosol. The strong affinity of the transferrin receptor for transferrin (its ligand) ensures that the cell will get all the iron it needs even if transferrin represents only a small fraction of the protein molecules present in the ECF. Receptor-mediated endocytosis is many thousand times more efficient than simple pinocytosis in enabling the cell to acquire the macromolecules it needs.

Another Example: the Low-Density Lipoprotein (LDL) Receptor

Cells take up cholesterol by receptor-mediated endocytosis. Cholesterol is an essential component of all cell membranes. Most cells can, as needed, either synthesize cholesterol or acquire it from the ECF. Human cells get much of their cholesterol from the liver and, if your diet is not strictly "100% cholesterol-free", by absorption from the intestine.

Cholesterol is a hydrophobic molecule and quite insoluble in water. Thus it cannot pass from the liver and/or the intestine to the cells simply dissolved in blood and ECF. Instead it is carried in tiny droplets of lipoprotein. The most abundant cholesterol carriers in humans are the low-density lipoproteins or LDLs.

LDL particles are spheres covered with a single layer of phospholipid molecules with their hydrophilic heads exposed to the watery fluid (e.g., blood) and their hydrophobic tails directed into the interior. Some 1,500 molecules of cholesterol (each bound to a fatty acid) occupy the hydrophobic interior of LDL particles. One molecule of a protein called apolipoprotein B (apoB) is exposed at the surface of each LDL particle.

The first step in acquiring LDL particles is for them to bind to LDL receptors exposed at the cell surface. These transmembrane proteins have a site that recognizes and binds to the apolipoprotein B on the surface of the LDL. The portion of the plasma membrane with bound LDL is internalized by endocytosis. A drop in the pH (from

5) causes the LDL to separate from its receptor. The vesicle then pinches apart into two smaller vesicles: one containing free LDLs the other containing now-empty receptors. The vesicle with the LDLs fuses with a lysosome to form a secondary lysosome. The enzymes of the lysosome then release free cholesterol into the cytosol. The vesicle with unoccupied receptors returns to and fuses with the plasma membrane, turning inside out as it does so (exocytosis). In this way the LDL receptors are returned to the cell surface for reuse.

People who inherit two defective (mutant) genes for the LDL receptor have receptors that function poorly or not at all. This creates excessively high levels of LDL in their blood and predisposes them to atherosclerosis and heart attacks. The ailment is called familial (because it is inherited) hypercholesterolemia.

Mutations in APOB, the apoB gene, cause another form of inherited hypercholesterolemia.

  • the retinoid vitamin A (retinol) bound to the retinol-binding protein
  • the steroids
      bound to the vitamin D binding protein bound to the corticosteroid binding globulin and estrogens bound to the sex hormone binding globulin
  • and there is growing evidence that, like cholesterol, they are taken into the cell by receptor-mediated endocytosis.


    Exocytosis

    Exocytosis is the process by which cells release particles from within the cell into the extracellular space.

    Learning Objectives

    Describe exocytosis and the processes used to release materials from the cell.

    Key Takeaways

    Key Points

    • Exocytosis is the opposite of endocytosis as it involves releasing materials from the cell.
    • Exocytosis has five stages, each leading up to the vesicle binding with the cell membrane.
    • Many bodily functions include the use of exocytosis, such as the release of neurotransmitters into the synaptic cleft and the release of enzymes into the blood.

    Key Terms

    • secretion: The act of secreting (producing and discharging) a substance, especially from a gland.
    • vesicle: A membrane-bound compartment found in a cell.

    Exocytosis

    Exocytosis’ main purpose is to expel material from the cell into the extracellular fluid this is the opposite of what occurs in endocytosis. In exocytosis, waste material is enveloped in a membrane and fuses with the interior of the plasma membrane. This fusion opens the membranous envelope on the exterior of the cell and the waste material is expelled into the extracellular space. Exocytosis is used continuously by plant and animal cells to excrete waste from the cells.

    Exocytosis: In exocytosis, vesicles containing substances fuse with the plasma membrane. The contents are then released to the exterior of the cell.

    Exocytosis is composed of five main stages. The first stage is called vesicle trafficking. This involves the steps required to move, over a significant distance, the vesicle containing the material that is to be disposed. The next stage that occurs is vesicle tethering, which links the vesicle to the cell membrane by biological material at half the diameter of a vesicle. Next, the vesicle’s membrane and the cell membrane connect and are held together in the vesicle docking step. This stage of exocytosis is then followed by vesicle priming, which includes all of the molecular rearrangements and protein and lipid modifications that take place after initial docking. In some cells, there is no priming. The final stage, vesicle fusion, involves the merging of the vesicle membrane with the target membrane. This results in the release of the unwanted materials into the space outside the cell.

    Some examples of cells releasing molecules via exocytosis include the secretion of proteins of the extracellular matrix and secretion of neurotransmitters into the synaptic cleft by synaptic vesicles. Some examples of cells using exocytosis include: the secretion of proteins like enzymes, peptide hormones and antibodies from different cells, the flipping of the plasma membrane, the placement of integral membrane proteins(IMPs) or proteins that are attached biologically to the cell, and the recycling of plasma membrane bound receptors(molecules on the cell membrane that intercept signals).


    The Back Story

    by Eleonora Aquilini

    What work led to this paper?

    Apicomplexan parasites are single-celled, obligate intracellular parasites defined by the presence of an “apical complex” - a group of cytoskeletal structures and organelles located at the anterior end of the cell. The apical complex is a critical compartment for parasites, because it regulates the events leading to host cell invasion. Out of all the remarkable structures of the apical complex, rhoptries are pear-shaped organelles that inject proteins into the host-cell (crossing not only the plasma membrane of the parasite but also that of the host), to support invasion and subversion of host immune function. But, how!?

    The molecular mechanism by which they are discharged and their effectors delivered into the host cytoplasm was unclear, and no orthologues of secretion genes from bacteria, yeasts, animals or plants were found associated to rhoptry secretion in Apicomplexa (with only one exception). Rhoptry secretion could thus represent an unconventional, parasite specific, secretory mechanism…

    Luckily, pioneering ultrastructural work done at the turn of the '80s pointed out some resemblance between secretory organelles in apicomplexan parasites and ciliates - their free-living relatives. Moreover, a rosette of 8 intramembranous particles, essential for fusion and secretion in free-living ciliates, was found to be present also in several apicomplexan parasites at the site of exocytosis, where the fusion takes place.

    A) Free-living Paramecium , Dubremetz et al. 1976 b) Eimeria parasite, Dubremetz et al. 1977 c) Plasmodium parasite merozoite, Dubremetz et al. 1979 d) Roof of Casa Batlo, by A. Gaudí in Barcelona – we were so obsessed with the rosette that we could spot it literally everywhere , Lebrun 2017 e) free-living Paramecium, Froissard et al. 2002 f) Plasmodium parasite sporozoite, Dubremetz et al. 1979 g) Toxoplasma parasite tachizoite, Aquilini et al. 2021

    These resemblances were all we needed to start.

    What did we do?

    Back and forth between free-living Ciliata and apicomplexan parasites.

    Building on the ultrastructural work done previously, we explored the similarities between organelle secretion system in ciliates (Tetrahymena thermophila and Paramecium tetraurelia) and apicomplexan parasites (Toxoplasma gondii and Plasmodium falciparum). We used cutting edge imaging techniques to study the structural elements and their architecture, and state-of-the-art molecular biology methods to explore the mechanism and molecular composition of these secretion systems.

    We cultivated tons of parasites and free-living ciliates, performed uncountables experiments, established the most amazing collaborations (Turkewitz Lab, specialists in secretory organelle biogenesis of Ciliata and for hobby in monarca butterfly metamorphosis and release and Chang Lab, masters of Cryo-electron tomography of the parasite apex in unfixed condition), I lived in Chicago for a while, we shared and discussed results in conferences (won one of the prizes at the Molecular Parasitology Meeting with preliminary results in 2018!), passed hours and hours and MORE hours looking at parasites under different types of microscopes . Dare I say we looked at them for hundreds of hours? Yes.

    What did we find?

    That rhoptry exocytosis depends on a fusion rosette of intramembrane particles visible on the plasma membrane at the site of secretion. Rosette formation requires the “non discharge” complex (Nd6, Nd9, NdP1, NdP2). This critical structure (the fusion rosette) and the genetic elements necessary for its assembly are akin to the exocytic machinery of ciliates, their free-living relatives. Our data suggests a common ancestry for this fusion machinery that adapted, in two groups of protists that diverged hundreds of millions of years ago, to radically different environments . This same machinery is involved in self- defence in the free-living ciliates and supports host-cell invasion in intracellular parasites.

    We also noticed something that caught our attention: one cardinal protein responsible for exocytosis was present on an enigmatic apical vesicle visible in at the very tip of several apicomplexan parasites, but a bsent in free-living ciliates. Our results indicate that this parasite-exclusive vesicle is sandwiched between the tip of the rhoptry and the rosette, and may reflect the additional complexity required for parasitism and cell invasion, in which exocytosis must be coupled with injection of rhoptry content through the host cell membrane – in contrast with free-living organisms, where secretory organelles are more simply discharged into the environment to thwart predators. In support of this hypothesis, a similar vesicle is present at the apex of Perkinsus marinus, an oyster’s parasite phylogenetically considered an ancestor both of free-living ciliates and apicomplexan parasites, and a key taxon for understanding unique adaptations to parasitism.

    CRYO-ET view of the fusion rosette of intramembranous particles visible on the plasma mambrane and sagittal view and 3D reconstruction of the interactions between the tip of the rhoptry (orange), the apical vesicle (pink) and the fusion rosette (purple).

    Next challenge? Understand precisely the role and constitution of the apical vesicle, and further clarify how the molecular elements interact with each other to promote membrane fusion and rhoptry exocytosis.

    ZOOM-party paper accepted! From left to right, top line: Maryse Lebrun, Eleonora Aquilini, Marta Cova in the middle: Yi-Wei Chang, Nicolas Dos Santos Pacheco, Aaron Turkewitz bottom line: Laurence Berry and Daniela Sparvoli

    Summary and Future Perspectives

    We have summarized the above-mentioned mechanisms regarding exocytosis, endocytosis and possible coupling factors in Figure 2. From a macroscopic view, exocytosis may be matched with endocytosis: full fusion with clathrin-mediated endocytosis, KR and KS with clathrin-independent endocytosis, and sequential fusion and multivesicular exocytosis with bulk endocytosis. In this sense, the fate of the components of the fusing vesicle may be pre-determined at the moment of its choice of fusion modes. Therefore, understanding the early fusion intermediates of a vesicle, such as the hemifusion state, pore opening, dilation, and shape retention, will be instrumental for the understanding of the whole coupled process.

    Figure 2. Different types of exocytosis, endocytosis and coupling factors in secretory cells. Coupling factors and their roles in different steps are also listed on the scheme.

    The listed classification of different exocytosis and endocytosis subtypes is not based on molecular mechanism but rather hinges on studies that involve different experiments conducted on different cell types. The terminologies defined by different methods may not be mutually inclusive or exclusive. For example, bulk endocytosis is usually regarded as a subcategory of clathrin-independent endocytosis. However, the bulk membrane invaginations observed in secretory cells under EM, which are often taken as evidence supporting bulk endocytosis, may support the internalization of small or large chunks of membrane in a clathrin-dependent manner in live cell studies. KR and KS may be one uniform process at different stages but could also be two distinct processes with non-overlapping mechanisms. To differentiate these controversies, it is important to sort out molecules that are exclusively used for some specific processes, in addition to actin for clathrin-independent endocytosis (He et al., 2008 Delvendahl et al., 2016). Alternatively, we shall examine the same process in the same cells using multiple techniques. For example, combining cell-attached membrane capacitance measurements with imaging vesicular lipids in endocrine cells will help clarify whether lipid exchange occurs between the vesicle and the plasma membrane during the flickering of a small fusion pore. Simultaneous imaging of vesicular components and extracellularly applied fluorescent dextran of different sizes will help monitor the dilation of a fusion pore from ߡ nm to a much larger in diameter (Takahashi et al., 2002). This will differentiate KR and KS and ultimately determine the size of fusion pores accompanying KS exocytosis. Monitoring the shape of the membrane may reveal clues of hemifusion in live cells (Zhao et al., 2016) and will also confirm or disapprove the compound fusion/multivesicular exocytosis theories and their physiological significance. Finally, operating at a nanometer scale with lifetimes of milliseconds, most of the fusion intermediate structures described here can hardly be directly discerned even with state-of-the-art super-resolution microscopy methodologies (Huang et al., 2009 Schermelleh et al., 2010). Despite differences in exocytosis kinetics and the organization of fusion sites between synapses and endocrine cells, we believe that the core exo-endocytosis coupling mechanism is conserved. Therefore, if we can improve the temporal and spatial resolution and duration of current super-resolution imaging technologies, direct visualization of fusion pore intermediates in endocrine cells may invoke new insights that would render much of the discussed theories here obsolete.


    Understand Cell Secretion in 4 Q&As

    In secretory cells, such as the secretory cells of endocrine glands, organelles related to the production, processing and “export” of substances are widely present and well-developed. These organelles are the rough endoplasmic reticulum and the Golgi apparatus.

    The nuclear membrane of secretory cells generally has more pores to allow the intense traffic of molecules related to protein synthesis between the cytoplasm and the nucleus.

    Secretory Organelles

    3. What is the role of the rough endoplasmic reticulum and the Golgi apparatus in the production and release of proteins?

    In its outer membrane, the rough endoplasmic reticulum contains numerous ribosomes, structures where the translation of messenger RNA and protein synthesis occur. These proteins are stored in the rough endoplasmic reticulum and are later moved to the Golgi apparatus. Within the Golgi apparatus, proteins are chemically transformed and, when ready, they are put inside vesicles that detach from the organelle. These vesicles fuse with the plasma membrane (exocytosis) in the right place and its content is released outside the cell.

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    Protein Degradation

    When a protein has outlived its usefulness or become damaged, it is degraded by the cell. In eukaryotes, a protein that is to be degraded has a number of copies of the small protein ubiquitin attached to it by a series of ubiquitin-adding enzymes. Ubiquitin serves as a tag that marks the protein for degradation. A tagged protein is then sucked into a large cellular machine called the proteasome, which itself is made up of a number of protein components and looks something like a trash can. Inside the proteasome, the tagged protein is digested into small peptide fragments that are released into the cytoplasm where they can be further digested into free amino acids by other proteases. The life of a protein begins in one cellular machine called the ribosome and ends in another called the proteasome.


    Watch the video: Cell Transport (January 2023).