ITPL Diary
The CoMPLEX ITPL programme is a crash course in the many
different ways we can measure what's going on in biological systems. In
a series of dense lectures, researchers co-opted from a variety of
fields describe techniques either developed or advanced to solve
problems in those fields. There's a lot of underlying theory (quantum
mechanics, photochemistry, physiology, etc), a lot of complicated
engineering (just to make the dratted processes happen), and then a lot
more difficulty in taking the outputs and turning them into something
that can perhaps go towards answering the real question, whatever
that may be. The whole mess can get a bit bewildering at times; which is
just how I like it :)
Anyway, following up on the lectures, a week was taken up by the ITPL
practicals, which is to say: spending each day at a different lab
getting variously exposed to a selection of those instrumental
techniques. And what a fun-filled experience it was.
Monday was spent in the basement of the Physics department, in a
room full of lasers1. The
particular techniques of concern were those of fluorescence lifetime
imaging, with a side order of anisotropy and energy
transfer.
A molecule fluoresces when one of its electrons is temporarily
excited to a higher energy level by an incident photon, only to drop
back to its ground state after an intermediate energetic
relaxation. The redistribution of energy in the interval results
in the emitted photon having a longer wavelength than the absorbed one
-- it is red-shifted -- and that difference is (statistically)
characteristic of the molecule2. The bit of molecule that does this is
called a fluorophore, and the time between it absorbing and
emitting is the fluorescence lifetime.
An ordinary microscope allows you to look at a reasonably small point on
your target, subject to various limitations3. If the target is illuminated with light
at the wavelength your fluorophore absorbs, the microscope will pick up
some of the corresponding emission photons, which will have a different
wavelength. Add in a filter to block all the photons that aren't that
wavelength and what comes out is your basic fluorescent image.
Now, suppose that the illumination occurs in very tiny pulses.
Fluorophores will be excited by a pulse and then -- sometime later --
emit. If you know exactly when you fired the pulse and you know exactly
when you picked up the emission, you can gauge the fluorescence
lifetime. Do that a lot of times and you'll get a reasonable picture of
the lifetimes at the point you're focussing on4.
If you repeat this process at different points across the sample, you
can build up an image of the fluorescence lifetimes in different
locations, which is a new layer of information on top of the
fluorescence intensity image you already have. Fluorescence
lifetimes are sensitive to the molecular environment, so this
information may be valuable.
Hey! Where are you going? Come back, there's more!
Fluorescence is orientation dependent. Molecules exist in space and can
point this way and that. The fluorophores have dipole
moments5 along which
their absorption characteristics manifest. The probability of an
incoming photon causing fluorescence is tied to its polarization. Angled
the wrong way, it's never gonna happen.
Since we control the polarization6 of the incident light and can also
filter the light we detect, it is possible to use this to estimate
orientational changes among the molecules over the fluorescence
lifetime. This is particularly interesting if we have (courtesy of
genetic engineering) a molecule with more than one fluorophore on it --
one end glows green, the other red -- because it can give information
about the relative orientations of the two fluorophores, and thus about
the conformation of the molecule. Which would take us on to
FRET, but this piece is already dead weight and I've only done
Monday...
Tuesday found me in Medical Physics as both experimenter and
guinea-pig.
Working out what's going on inside people's bodies -- and heads -- is a
problem of pretty much unlimited scope. Bodies are complicated things
and their owners have attitudes. They don't like being prodded and
probed7. They get litigious
when things go wrong or, you know, whenever they just feel like it. They
are, basically, a pain in the arse.
They're also pretty fragile. You can't just go tearing in there to see
what's going on, there'll be blood and guts and all sorts of mess. It'll
end in tears.
As far as possible, you want to look inside people non-invasively. If
only they were transparent.
Well, that may be too much to ask, but it turns out that many human body
tissues are sort of translucent. We occlude, but not completely.
There are wavelengths of light that pass through us relatively easily.
Some, like X-rays, are horribly energetic and damaging, only to be used
as a last resort. But the near infra-red region also turns out to be
handily penetrative without all those nasty ionizing side effects.
It's not transparency -- IR beams don't pass neatly through the body to
be focussed into a crisp image on the far side -- but flesh is permeable
enough to IR to allow all kinds of measurements to be made. The rays are
scattered, but only partially absorbed -- and the degree to which
they're absorbed can reveal interesting things, such as the oxygenation
level of the blood8.
While this is medically useful already -- and you've quite likely had
the measurement taken yourself, you can do it in Boots these days -- for
our purposes it's just a stepping stone towards more detailed
information. The body needs oxygen in order to do work and is generally
quite good at getting it to where that work is happening. So measuring
local oxygenation in different places can give a good functional
view of where the action is.
Which is why I wound up with optodes all over my head doing anagrams in
a darkened room.
Actually constructing a useful image from such measurements --
and even just making the measurements themselves -- is a painstaking
business because of the low level of transmission, the high levels of
noise, various confounding physiological processes and the awkwardly
complex mixture of stuff we contain. The resolution is never going to be
high, but it's safe and practical and potentially important.
Wednesday was spent at the Cell Biophysics lab at Cancer Research
UK, down by Lincoln's Inn, doing some sample preparation and mass
spectrometry. The samples in question were membrane lipids from sea
urchin gametes, and the preparation involved many stages of dissolving
and filtering and centrifuging and drying and such: proper biochemistry
with test tubes and everything. The mass spec was much duller by
comparison, very impressive machinery but mostly computer-controlled and
rather slow.
Phospholipids are fatty molecules that make up the bulk of cell
membranes, key structural components of all lifeforms more complex than
a virus. Membranes were traditionally viewed as rather simple things,
thin waterproof bilayers held together by hydrophobia9, almost trivial beside the mind-boggling
complexity of DNA and proteins. However, given the enormous range of
interactions between the inside of a cell and the big bad world, and the
astonishingly precise control most cells can exercise over what goes in
and out, it is clear that membranes are actually seething hotbeds of
vital activity. While much of that is mediated by embedded proteins, the
lipid composition is also intricately involved. Determining that
composition in detail is no simple task.
Mass spectrometry, which uses an electric field to separate out
particles according to their mass-charge ratio, can help, provided the
particles are actually charged. To get it to work, the lipids are broken
into fragments and the resulting spectrum analysed to see which
fragments came from where. Of course, there are a lot of different
possible lipids with varying arrangements of fatty acid chains, fragment
masses and charges. With only a single spectrum there would be too much
noise to get much sense of where the spikes were, so the feed into the
mass spec is first run through a high-pressure liquid chromatography
column, which spreads the fragments out according to their
hydrophobicity.
The resulting data set is still pretty intimidating. I'm very glad not
to be responsible for analysis beyond a casual conversation about a
single experiment. Our mentor for the day was dealing formally with
hundreds or maybe thousands of them...10
Thursday was back to UCL for some very hands-on patch
clamping in the Pharmacology.
Among their many exciting features, lipid membranes can contain little
protein assemblies called ion channels that control the movement
in and out of particular ions. These nanomechanical marvels can
discriminate11 the types
of individual atoms and if they're not on the guest list they just don't
get through. That's atoms we're talking about. Do you have
any idea how small they are?
The passage of specific ions through the ion channels constitutes an
electric current, and handling such currents is one of the things cells
do that makes them alive. The currents are not large -- of the
order of pico-amps -- but they're important to the cell and also
-- rather amazingly -- measurable in the lab.
In order to do that, you need to isolate some channels on the cell
surface with a very tiny glass tube: both the glass and the cell
membrane are good insulators, and when they're firmly clamped together
the only route left through which current can flow is the ion channels.
First catch your tiny glass tube.
Creating a microscopic pipette for patch-clamping is a surprisingly
simple, surprisingly manual and also somewhat hit and miss
business. Basically, you take a small glass tube, heat it in the middle
until it gets soft, and then pull the ends apart until it breaks. There
are a bunch of refinements along the way, but given favourable
circumstances that process will give you the pipette you need; on top of
which it's also rather fun.
You then fill it with electrolyte and attach it to an electrode on a rig
that can gear down simple hand cranks to produce very sensitive
adjustments in each direction. The whole caboodle is attached to a
powerful microscope: you carefully maneouvre it down onto a cell et
voilà. There's a giga-ohm resistance seal and the only
currents that can flow are through the ion channels.
It's not quite as haphazard as I paint it, but you do have to
take a certain amount on trust. Which may be why it's so gratifying when
it works.
Ion channels are far too small to see with your own eyes, no matter how
powerful the microscope, thanks to the resolution limit mentioned in footnote 3. So there's no telling what's included in the
patch of membrane to which your pipette is attached. But as you increase
the membrane potential -- which is one of the things that can tell the
ion channels to open -- you start seeing a tiny current flow, increasing
or decreasing in very obvious steps, all the same size.
A single ion channel lets through a fixed amount of current when it's
open; all channels of the same type having the same conductance. So each
of those steps represents a single open channel. What you're "seeing" as
the current jumps up and down is the opening and closing of those
channels -- something so small it is barely possible to imagine.
How cool is that?
Finally, on Friday I was in Physiology, somewhere in the depths
of the medical school. The experiment combined both the
electrophysiological patch clamping of the day before with fluorescence
techniques related to Monday's, only this time it was two-photon
fluorescence. Used, in this case, to observe a living neuron in
something approaching 3d.
In ordinary one-photon fluorescence, as described earlier, you shine
energetic (usually UV) light through your sample and everywhere it hits
a fluorophore some less energetic light is emitted. This is a great
process, but it has disadvantages. All that UV light pouring in can
damage the target tissues. Fluorophores are often unstable and can stop
being responsive after even relatively short exposures. Since any
fluorophore in the light's path will fluoresce, not only can you
photobleach what you're looking at but everything nearby too. Worse, you
have to jump through some vexing optical hoops to eliminate all the
emissions that aren't coming from your focal point, and in the process
you sacrifice a lot of photons that you can't afford to lose.
In the two-photon technique, the fluorescence is triggered by two
lower-energy photons hitting the fluorophore at the same time.
That may seem obvious, but there's a problem: the ground state and the
excited state are strict quantum energy levels. There is nothing
in between. Really, it should be impossible to get bumped from
one to the other in two stages.
Enter Heisenberg's Uncertainty Principle, which states that there
is an innate vagueness in the energy of a system in a certain amount of
time, and vice versa. This uncertainty is very, very small: even
for the really tiny energy state differences we're talking about in our
fluorophore, the margin for error is minuscule. But it does exist. As
long as the two photons hit the fluorophore virtually
simultaneously, they can be absorbed and everyone's happy.
In order for there to be any chance of it happening at all, there have
to be a vast number of photons in the immediate vicinity of the
fluorophore, which sounds like a nuisance but is actually the greatest
benefit of two-photon fluorescence: it only happens where the photon
flux is highest, which is exactly at the focal point.
The rest of your sample doesn't fluoresce!
You don't need to worry about filtering emissions from everywhere else,
because there aren't any. The lower energy light you're shining into
your sample just blunders harmlessly through. Every higher-energy
emission photon you collect has to be from the point you're looking at.
You can scan a live cell in rastered layers and build up a complete 3d
picture of the fluorophores within it.
Of course, it's a delicate procedure and you'll need hundreds of
thousands of pounds worth of equipment to do it, but at least it's
easier than three-photon fluorescence...
1 Cover your eyes if you bend down to pick something up.
2 This is the process that makes dayglo items show up under the ultraviolet lights in nightclubs -- they absorb the high energy UV photons and emit lower energy visible ones. The loss goes, pretty much, to heat, though this effect is vanishingly small compared to that of you shaking your funky groove thang.
3 Probably the most important limitation is the wavelength of light you're viewing with. Imagine trying to measure a grain of sand with a yardstick marked in inches -- there just isn't enough resolution to make a meaningful measurement. Light is a yardstick with much finer markings than that, but there's still space between the markings and that space limits what you can resolve.
4 Naturally, things are not quite so simple. You're dealing with molecules and photons en masse. A single excitation event will (perhaps) lead to a single emission event after a particular time, and the emitted photon may or may not head towards your detector. Other events will have other lifetimes and go off in other directions. What you detect is a fraction of the available events with a range of different lifetimes. So the result is not a single lifetime, but a distribution of them.
5 Don't even ask, okay?
6 We control the vertical. We control the horizontal.
7 Oh come on. Just insert your own smutty innuendo here.
8 Another simplification, of course. You need to take measurements at more than one wavelength and do a bit of linear algebra, but the details are relatively straightforward and the basic technique has been around since WW2.
9 Not, in this case, a synonym for rabies; rather an expression of the tendency of some molecules or parts thereof to interact unfavourably with water, a substance overwhelmingly present throughout all biological systems.
10 That's a PhD for you. In a year's time I may be facing something similar...
11 Let's just ignore the implicit anthropomorphism of this sentence.
1 Cover your eyes if you bend down to pick something up.
2 This is the process that makes dayglo items show up under the ultraviolet lights in nightclubs -- they absorb the high energy UV photons and emit lower energy visible ones. The loss goes, pretty much, to heat, though this effect is vanishingly small compared to that of you shaking your funky groove thang.
3 Probably the most important limitation is the wavelength of light you're viewing with. Imagine trying to measure a grain of sand with a yardstick marked in inches -- there just isn't enough resolution to make a meaningful measurement. Light is a yardstick with much finer markings than that, but there's still space between the markings and that space limits what you can resolve.
4 Naturally, things are not quite so simple. You're dealing with molecules and photons en masse. A single excitation event will (perhaps) lead to a single emission event after a particular time, and the emitted photon may or may not head towards your detector. Other events will have other lifetimes and go off in other directions. What you detect is a fraction of the available events with a range of different lifetimes. So the result is not a single lifetime, but a distribution of them.
5 Don't even ask, okay?
6 We control the vertical. We control the horizontal.
7 Oh come on. Just insert your own smutty innuendo here.
8 Another simplification, of course. You need to take measurements at more than one wavelength and do a bit of linear algebra, but the details are relatively straightforward and the basic technique has been around since WW2.
9 Not, in this case, a synonym for rabies; rather an expression of the tendency of some molecules or parts thereof to interact unfavourably with water, a substance overwhelmingly present throughout all biological systems.
10 That's a PhD for you. In a year's time I may be facing something similar...
11 Let's just ignore the implicit anthropomorphism of this sentence.