A A physics perspective needed on how the Mitochondria machine works

I purposely added this post to the general physics thread rather than the biology thread because the solution is probably not in biology.

Basically my PhD is in mitochondrial biology. Mitochondria are the energy producing organelles in the body which make energy by adding a chemical bond converting a di-phosphate ADP to ATP (tri-phosphate).
The problem is the precise machinery about how this works is not entirely known. The oxidative phosphorylation system which is composed of 5 large protein complexes known as complexes I-V are responsible for making ATP which is shown in any high school textbook. These mitochondria have double membranes and can be isolated from cells and tissues and treated like individual cells on their own.

However, in the past 20 years, through the use of less volatile detergents used for breaking the mitochondrial membrane, we have learned that these individual complexes can combine together in a dynamic way under different metabolic cues to fine tune the system and make mitochondria more efficient. This happens in exercise for instance.

But the act of breaking the membrane may shatter this protein machine into different 'parts' and the parts are what we see. These parts are called supercomplexes.
So how could we see these protein-protein interactions intact? People used formaldyhyde to cross link proteins and using this method found what they called the megacomplex, larger than supercomplexes.
But I think that these complexes all work together in a respiratory string, which makes ATP but we just can't see it.

I was considering using antibodies to view these complexes however antibodies are 1500 Da while proteins 100 times smaller in size are the ones which penetrate the membrane. So the act of using antibodies would breach the membrane.

Maybe the use of a hypotonic solution in which mitochondria uptake into their core might work. Then we could spin the isolated mitochondria down, transfer them into another solution and trigger the solidification of the uptaken hypotonic solution which would leave an imprint of proteins on the inner mitochondrial membrane and viewed using a microscope such as an electron-microsope?

It would be one of the most important questions to answer in biology, how does this machine work. But I don't think the answer is in biology. Perhaps there is some tool in physics or engineering that people routinely used and could be applied to this question. I need a non-biologist perspective. I appreciate your input.
 
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Maybe the use of a hypotonic solution in which mitochondria uptake into their core might work. Then we could spin the isolated mitochondria down, transfer them into another solution and trigger the solidification of the uptaken hypotonic solution which would leave an imprint of proteins on the inner mitochondrial membrane and viewed using a microscope such as an electron-microsope?

It would be one of the most important questions to answer in biology, how does this machine work. But I don't think the answer is in biology. Perhaps there is some tool in physics or engineering that people routinely used and could be applied to this question. I need a non-biologist perspective. I appreciate your input.
The solidification (cross-linking or freezing) of hypotonic solvent to prepare electron microscopy substrate is likely to denaturate protein complex. To examine the shape of such delicate structure, AFM (atomic force microscopy) would be less damaging and require less preparation than electron microscopy.
 
The solidification (cross-linking or freezing) of hypotonic solvent to prepare electron microscopy substrate is likely to denaturate protein complex. To examine the shape of such delicate structure, AFM (atomic force microscopy) would be less damaging and require less preparation than electron microscopy.
Ok, but the problem of any microscopy technique that I know of is that it is unable to examine the structure of these complexes from the inside-out of these organelles. So have to come up with a way to visualize these complexes while penetrating a double membrane structure to see them in their natural state.

The methods currently employed include lysing the mitochondrial membrane and using cryogenic electron microscopy to look at the shape of these proteins. We can model megacomplexes and propose the existence of a respiratory string but we do not have the ability to see them yet because we have to lyse the mitochondrial membrane.
It requires a radical new technique to investigate this question.

https://www.cell.com/cell/fulltext/S0092-8674(17)30887-5?_returnURL=https://linkinghub.elsevier.com/retrieve/pii/S0092867417308875?showall=true
 
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Ygggdrasil

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Biophysicists have developed light microscopy techniques, collectively called superresolution microscopy, that are capable of imaging at around 10-20 nm resolution, which may enable you to see these structures in live cells. In particular, researchers have developed a technique called tcPALM that has been used to see the transient formation of clusters of RNA polymerase molecules that form at a gene during a transcriptional burst (http://science.sciencemag.org/content/341/6146/664.long). These clusters are very short lived (~5s), so the technique is capable of seeing fairly dynamic interactions. Of course, the technique was looking at only homotypic interactions, and heterotypic interactions would be harder to monitor since they would require aligning multiple different color channels on the microscope. Multi-color single molecule tracking methods can also help visualize transient interactions between proteins diffusing in membranes (https://www.cell.com/biophysj/fulltext/S0006-3495(15)03214-2).
 
Biophysicists have developed light microscopy techniques, collectively called superresolution microscopy, that are capable of imaging at around 10-20 nm resolution, which may enable you to see these structures in live cells. In particular, researchers have developed a technique called tcPALM that has been used to see the transient formation of clusters of RNA polymerase molecules that form at a gene during a transcriptional burst (http://science.sciencemag.org/content/341/6146/664.long). These clusters are very short lived (~5s), so the technique is capable of seeing fairly dynamic interactions. Of course, the technique was looking at only homotypic interactions, and heterotypic interactions would be harder to monitor since they would require aligning multiple different color channels on the microscope. Multi-color single molecule tracking methods can also help visualize transient interactions between proteins diffusing in membranes (https://www.cell.com/biophysj/fulltext/S0006-3495(15)03214-2).

So I do have access to super-resolution microscopy, we used super-resolution and 3D electron microscopy to discover the existence of mitochondrial nanotunnels, however the main issue would be penetrating the double membrane of the mitochondria. I think dendra-2 may be too large to get inside mitochondria and used as a tag. In fact most tags, like GFP for instance would be too large to penetrate the membrane. That is the main problem, so have to think of a way to either get a tag inside without damaging the membrane or use another technique altogether. People have used FRET to look at interactions between complexes but it can't inform you on the existence of the respiratory string.
 

Andy Resnick

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we have learned that these individual complexes can combine together in a dynamic way under different metabolic cues to fine tune the system and make mitochondria more efficient.
This is an interesting problem, not sure I have much to offer. My first thought was to try patch clamping, for example:

https://www.nature.com/articles/330498a0

but that thought is based on an assumption that may not be valid. So, let me ask some clarifying questions:

1) What are the lengthscales and timescales that are most relevant?
2) How do you determine 'combine together'? Imaging is one way, but functional assays are another approach that can work.
3) What parameters do you use to define 'fine tune' or 'efficient'?
4) Can you make any simplifying assumptions, such as: the mitochondrial membrane proteins are distributed homogeneously, as opposed to spatial variations?
5) Has any (correlative or causative) relationship between a particular 'metabolic cue' and a particular combination been observed?

I guess that's enough for now.... like I said, it's an interesting problem!
 

Ygggdrasil

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So I do have access to super-resolution microscopy, we used super-resolution and 3D electron microscopy to discover the existence of mitochondrial nanotunnels, however the main issue would be penetrating the double membrane of the mitochondria. I think dendra-2 may be too large to get inside mitochondria and used as a tag. In fact most tags, like GFP for instance would be too large to penetrate the membrane. That is the main problem, so have to think of a way to either get a tag inside without damaging the membrane or use another technique altogether. People have used FRET to look at interactions between complexes but it can't inform you on the existence of the respiratory string.
I'm not sure what you mean. Fluorescent protein tags like dendra2 or GFP are genetically encoded and would be delivered to the mitochondria by the native mitochondrial import machinery with the tagged protein. Are you saying that proteins tagged with FPs are not trafficked correctly to mitochondria?
 
This is an interesting problem, not sure I have much to offer. My first thought was to try patch clamping, for example:

https://www.nature.com/articles/330498a0

but that thought is based on an assumption that may not be valid. So, let me ask some clarifying questions:

1) What are the lengthscales and timescales that are most relevant?
2) How do you determine 'combine together'? Imaging is one way, but functional assays are another approach that can work.
3) What parameters do you use to define 'fine tune' or 'efficient'?
4) Can you make any simplifying assumptions, such as: the mitochondrial membrane proteins are distributed homogeneously, as opposed to spatial variations?
5) Has any (correlative or causative) relationship between a particular 'metabolic cue' and a particular combination been observed?

I guess that's enough for now.... like I said, it's an interesting problem!
So the different parts 'supercomplexes' appear to assemble at different times. Some supercomplexes have been shown through pulse-chase to assemble as soon as 0.5 hrs but others do not assemble for many hours later. So it will be on the order of hours.
The tools for investigating this question are extremely limited, what people typically do is lyse the membrane and pass the native complexes through a Blue Native Gel, and they are later detected using immunoblotting. You can use a 2D gel gel to look at the structure and see which complexes localise together.
All we have now are correlations, when we treat animals under certain diets or if they undergo exercise, we see a change in the distribution of these supercomplexes. Its still a hypothesis that these supercomplexes lead to more efficienct mitochondrial systems but the evidence is mounting. I am looking at this system right now and its output.
I have analysed mitochondrial bioenergetics indirectly by quantifying the level of oxygen consumption used to generate ATP over time in response to different inhibitors which basically can inhibit specific complexes in this system by using a seahorse machine from agilent. Our drugs enhanced mitochondrial bioenergetics and I have correlated it with changes in supercomplex assembly, but thats another project.
 
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I'm not sure what you mean. Fluorescent protein tags like dendra2 or GFP are genetically encoded and would be delivered to the mitochondria by the native mitochondrial import machinery with the tagged protein. Are you saying that proteins tagged with FPs are not trafficked correctly to mitochondria?
They are trafficked to the mitochondrion, however as far as I know these tags are not able to incorporate into the matrix for instance. When you attach a GFP to lets say, a subunit of the oxidative phosphorylation system with a mitochondrial targeting sequence, the GFP is targeted to mitochondria but essentially gets stuck in the intermembrane space because of its size as it tries to shuttle through the mitochondrial pores. The problem will be at the mitochondrial import machinery.So you can use these tags to image the morphology of mitochondria in a cell or as a tool for mitochondrial localisation, but I haven't see tags used to distinguish interactions between proteins inside mitochondria for this reason. I would be glad if you could prove me wrong and show someone doing this because I may be able to incorporate their system.

Thats why I was thinking that the traditional route of flourescent tagging and imaging will not work. Someone else would have done it by now if it were that easy.
 
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This article is (tiresomely) walled off from profanum vulgus:

10.1055-s-0033-1340639-1.jpg


PF's xenforo implementation in this instance insists on copying to PF's servers the image that was specified inside bbs [img-/img] tags and then presenting an irretrievably shrunken version of it inside its own generated [attach-/attach] tags; here's the unadulterated authorized preview link if you want to see a less low-res version:
https://www.thieme-connect.de/media/synlett/201408/lookinside/10.1055-s-0033-1340639-1.jpg

And this one is available to the unwashed masses, and although it shows off a woefully weak steganangraphic/crrytographic/password based scheme for improvement over use of plaintext invisible ink messages, it also presents some intriguing ideas suggesting possibilities for micro-scale bio-investigative techniques: https://www.nature.com/articles/ncomms11374.pdf
 

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Ygggdrasil

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They are trafficked to the mitochondrion, however as far as I know these tags are not able to incorporate into the matrix for instance. When you attach a GFP to lets say, a subunit of the oxidative phosphorylation system with a mitochondrial targeting sequence, the GFP is targeted to mitochondria but essentially gets stuck in the intermembrane space because of its size as it tries to shuttle through the mitochondrial pores. The problem will be at the mitochondrial import machinery.So you can use these tags to image the morphology of mitochondria in a cell or as a tool for mitochondrial localisation, but I haven't see tags used to distinguish interactions between proteins inside mitochondria for this reason. I would be glad if you could prove me wrong and show someone doing this because I may be able to incorporate their system.

Thats why I was thinking that the traditional route of flourescent tagging and imaging will not work. Someone else would have done it by now if it were that easy.
It seems like FPs should be able to get into the mtiochondrial matrix. For example, here's a paper where the authors are able to get GFP into the mitochondrial matrix by tagging with the targeting presequence from COX8: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2141758/
 
It seems like FPs should be able to get into the mtiochondrial matrix. For example, here's a paper where the authors are able to get GFP into the mitochondrial matrix by tagging with the targeting presequence from COX8: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2141758/
Yes, I did this experiment 10 years ago in neurons. You see the diffusion of photoactivatable GFP through the mitochondrial network as a proxy for the rate of fusion. But if you read the paper, in order to get into the matrix, the GFP will be unconjugated, it will no longer be bound to COX8 inside the mitochondria because it will be cleaved by the presequence pepsidase or something similar, essentially making the protein small enough to get inside I suppose. Most nuclear proteins have their mitochondrial sequence cleaved for import so I guess the plasmid was designed to have the entire COX region cleaved off. I haven't looked at the plasmid map. So you have free floating GFP not targeted to anything particularly within the matrix. You couldn't use the technique to target specific mitochondrial proteins inside because to do that, you need the GFP conjugated, which essentially makes the protein a whole lot bigger and it gets stuck in the mitochondrial pores. At least thats my take on the paper unless I misread. Plus you also have the problem of a conjugated GFP tag if you did manage to get it inside the matrix not incorporating into the protein complex because of spacial issues and disrupting the stoichiometry. So I wouldn't be able to use it to address my question of super complex arrangement for both those reasons.
I wonder if there would be way to introduce a GFP tag inside and then somehow have it bind to a protein pocket or something associated with these complexes.
 
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Ygggdrasil

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I wonder if there would be way to introduce a GFP tag inside and then somehow have it bind to a protein pocket or something associated with these complexes.
Yes, this is possible using a split gfp system (https://www.nature.com/articles/nbt1044). You can tag your protein of interest with a 16 aa tag consisting of the 11th beta strand of GFP. If you import the rest of GFP into the matrix, it can bind to the 11th strand to reconstitute the full fluorescent GFP molecule. This system has also recently been adapted for superrresolution microscopy (https://www.nature.com/articles/s41467-017-00494-8).

Your point about the effects of GFP or other protein tags on complex formation, however, are still valid. Many FP variants are prone to aggregation and can alter complex stoichiometry (https://www.nature.com/articles/nmeth.1955), so careful choice of monomeric variants and validation of their effects on complex formation would be required.
 

Andy Resnick

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So the different parts 'supercomplexes' appear to assemble at different times. Some supercomplexes have been shown through pulse-chase to assemble as soon as 0.5 hrs but others do not assemble for many hours later. So it will be on the order of hours.
The tools for investigating this question are extremely limited, what people typically do is lyse the membrane and pass the native complexes through a Blue Native Gel, and they are later detected using immunoblotting. You can use a 2D gel gel to look at the structure and see which complexes localise together.
All we have now are correlations, when we treat animals under certain diets or if they undergo exercise, we see a change in the distribution of these supercomplexes. Its still a hypothesis that these supercomplexes lead to more efficienct mitochondrial systems but the evidence is mounting. I am looking at this system right now and its output.
I have analysed mitochondrial bioenergetics indirectly by quantifying the level of oxygen consumption used to generate ATP over time in response to different inhibitors which basically can inhibit specific complexes in this system by using a seahorse machine from agilent. Our drugs enhanced mitochondrial bioenergetics and I have correlated it with changes in supercomplex assembly, but thats another project.
Blargh... well, nothing I could think of would work. A few thoughts for you to consider:

1) The way I understand what you wrote, your project is taking results obtained from whole-animal studies (diet and exercise) and going directly to single molecule methods. Granted, you are working with cell cultures on the Seahorse, but even so, trying to go directly to single-molecule imaging (light or electron) seems like tossing a lit match into a wet room and hoping something will catch fire. But hey- maybe freeze-fracture will work.....
2) Your system of interest (I'll call it the Mitochondrial Interactome) seems to preclude most existing methods used to demonstrate protein-protein interaction: yeast 2-hybrids, FRET, protein arrays, etc. won't work either because you are interested in (I assume) transmembrane proteins or because you are looking specifically at proteins located within the mitochondria- just wondering, are these proteins coded by mtDNA?
3) I'm still going to suggest patch-clamping, especially when used with ion-specific electrodes. I think, for example, a measurement of the reversal potential could provide some insight.

Honestly, I think you (and your advisor) are going to have to invent the method. Then you can use it to answer your question.

Good luck, and let us know how it goes!
 

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