Nikon Eclipse Ti back port to Back Focal Plane distance specification needed

[Yossi_Steinberger]

Hello all,

I am designing a single-molecule microscope using a Nikon Eclipse Ti-E inverted microscope. My setup involves focusing a collimated laser beam onto the back focal plane of a 100×/1.4 NA objective via a ~200mm external lens mounted just outside the microscope’s back port (photo of the Thorlabs adapter attached).

Does anyone know the optical path distance from the microscope’s back port to the objective’s back focal plane? I haven’t found this specification in the product documentation.

Thank you,
Yossi

[P_Tadrous]

Hello. I don’t know much about Nikon specifically but it is highly unusual for objectives to have their BFP outside the objective at all (except for some very low magnification objectives like some x4 and below - but even then not always depending on the exact model).

There have been some microscopes specifically made in the past that allow access to a conjugate of the objectives’ BFP in a standardised position up the tube but those are very specific models of scope which were designed for direct access Fourier filtration with a SLM.

I note from Nikon’s brochure that the Ti2-E has the ability to do ‘external’ phase contrast with any (non-phase) objective by means of a phase ring in the eyepiece base unit which implies that that model provides access to a conjugate of the BFP in a standardised position in the eyepiece base unit. Not sure about the ordinary Ti model though.

PJT

[Yossi_Steinberger]

Hello @P_Tadrous

Thanks for the reply—and I realize my original question may not have been clear.

I’m not trying to access the back focal plane (BFP) outside the objective. I’m using the rear port of a Nikon Eclipse Ti (non‑Ti2) and plan to place a focusing lens just outside that port to focus a collimated laser onto the objective’s BFP (Nikon 100×/1.4 NA oil, HP Plan Apo VC).

What I’m looking for is the nominal optical distance from the rear‑port reference surface to the objective’s BFP along the epi‑illumination path. If this depends on configuration (e.g., filter cube, magnification changer), a typical value, a Nikon drawing/spec, or a practical way to measure it would be extremely helpful.

In a previous post of mine, the value 200mm was suggested, but not with certainty.

Coupling lasers to a widefield microscope?

Dear μforum - I have a Zeiss Axioobserver microscope, and a EMCCD camera, and wish to purchase a laser light source, and to set up a single molecule imaging system in my lab (I am a PhD student). However, I am ignorant of a good way to couple the lasers to the microscope. In particular, I want to use the 100x, 1.4 numerical aperture objective, and to achieve a maximum power of 30 kW/cm^2 with a far-red laser of ~640nm, but to achieve much lower powers of 30 W/cm^2 with a 405nm, 488nm and 549nm…

Many thanks,
Yossi

[P_Tadrous]

Hopefully someone of specific knowledge of your microscope can answer this. In the mean time, again speaking generically, it is unusual for the BFP of a set of objectives to lie in a single plane unless the objectives are specially designed as a set to do this for some reason. So, if you plan to use more than one objective for your experiment you will need to know the position for each objective separately and adjust the distance whenever you change objectives.

To measure the distance there are a few methods. You can shine a parallel beam up from the front of the objective and measure where it comes to a point out the back - although that would be impractical because most objectives have their BFP inside their barrel, esp. high power ones.

Another method is to focus a tiny spot of light onto the centre of the back of the objective and move the spot’s position (in length) till a parallel beam comes out the front of the objective - i.e. empirical trial and error. There is no need to use a laser for this but the spot must be tiny or you will never get a parallel beam. Practically, because of the difficulty in getting a tiny spot and because of the dangers of using lasers, it might actually be more practical to just focus an image of something (like an EM grid mask) on the back of the objective and place a converging lens of known focal length in front of the objective. Then measure the light distribution at the focal length of the lens you placed in front of the objective. When that light distribution is in the form of a focussed image of your mask you know that the primary mask image is focussed at the BFP of the objective.

Make sure that any mag changers or filter cubes you intend to use in your experiment are also in place when you do this. This is probably the most practical approach for you - unless someone else here can give you more specific guidance to your model.

PJT

[Omnistic]

@Yossi_Steinberger

I’ve done exactly what you did on a Nikon Eclipse Ti2, and to my surprise the tube lens (your 200mm lens) had to be quite inside the microscope body. Its not exactly like this anymore, but this is a screenshot from my original design.

You can see the THORLABS SM2N1 adapter and the 200-mm lens in blue. In the next screenshot you have a rough idea of the distance: ~115mm.

But keep in mind that I got there by trial and error. I couldn’t find the actual information. Therefore, I used the rods be able to translate the lens along the optical axis and properly align it.

The next question is how should you align the lens then? Because this alignment wasn’t crucial for me, this is how I approached this problem.

DISCLAIMER: this is a dangerous procedure and should use laser safety equipment, I take no responsibility for the content of this post and if you are following the instructions below you will do so at your own risk.

You know that if your beam is focused in the back focal plane of the objective lens, it should come out collimated after the objective lens. You kind of want to avoid using the immersion lens because the following measurement would be harder to setup. Change to a low magnification objective (or at least one without immersion) and assume the parfocality is preserved by changing the lens. This assumption holds better if the lenses are from the same manufacturer. Then, shine your laser through the back port. Tilt the condenser column to get it out of the way and let the laser beam reach the ceiling. Move the 200-mm lens along the optical axis until you make the smallest possible spot on the ceiling. In doing so, you’ll focus the beam at “infinity” (the higher the ceiling the better), thereby collimating it.

As a side note, a free space collimated laser beam is quite dangerous because the light intensity remains approximately constant as it travels.

Next time I realign my setup, I’ll take some pictures and measurements for you. But I hope this still helps.

Take care,

Omni

[Yossi_Steinberger]

Hello @ Omnistic

Can you please describe your system’s initial light source, collimator, and additional downstream optical elements, and their positions?
It looks like you are using kohler illumination? Or is it critical illumination with a relay?

I would like to have a 200 um or 400 um NA 0.22 multimode fiber nearly fill, ie, just underfill, the back focal plane of my 100X NA 1.4 objective, such that I have a widefield illumination field somewhere between 40-80 um in diameter. I am stuck with how to go from the multimode fiber output to the back focal plane.

Thank you for the excellently visualized setup.
Yossi

[Omnistic]

@Yossi_Steinberger

For general reference, the work that I did was to implement the setup from this paper in our Ti2.

Regarding your questions, I am of course happy to describe what I did in more details. However, I am not totally sure what exactly you are trying to achieve. Perhaps a drawing would help.

In your original post you wrote:

focusing a collimated laser beam onto the back focal plane

Is it still what you are trying to do here?

[Yossi_Steinberger]

Hello @ Omnistic

Thank you for the quick response.

“Is it still what you are trying to do here?” Yes. More specifically:

Category	Detail
Biological Context	Single protein and single genome position in live yeast cell.
Protein Labeling	Fused to HaloTag, used with JFX650 dye (appears as red spot).
Genome Position Labeling	Marked with LacO system, LacI fused to mNeonGreen (appears as green spot).
Nuclear Membrane Marker	mTagBFP2.
Imaging Exposure Times	Protein: 5ms (ideal 500us); DNA: ~20ms; Nuclear Membrane: 20ms or longer.
Illumination Requirements	JFX650 needs ~3 kW/cm² for 5ms exposure.
Camera/ROI Strategy	Prefer smaller region of interest and smaller illumination field due to row readout time (~10us/row).
Desired Illumination Field	~70 µm diameter.
Light Source	400µm diameter 0.22 NA multimode fiber.
Lasers	30 mW 405 nm diode, 30 mW 488 nm diode, 1 W 640 nm diode. All free running, Fabry Perot. Despeckled.
Illumination Method	Critical epi illumination with relay (not Kohler).
Optical Setup	12.5 mm collimator (L1), 200 mm L2, 180 mm L3.
Objective	100x 1.4 NA. Back focal plane aperture ~5.6 mm; beam ~4.95 mm.
Lens Placement	L3 inside Nikon Ti2-E microscope body.
Focus Adjustment	System on rail for axial movement to find correct focus.

If this setup could work, how would I obtain the appropriate L3 lens to back focal plane optical path distance?

Thank you,
Yossi

p.s., I heavily used LLMs to inform the design and also the visualization below.

[Omnistic]

@Yossi_Steinberger

Unfortunately, I am still not understanding what you are trying to do. Keep in mind, I am a modest optical engineer with limited time to help (for free).

With that in mind, I’ll give you a few tips and recommendation and perhaps that can get you started.

When we (optical engineers) place optical components we either have control over the optomechanics and we can insure accurate placement by design. Or we rely on alignment. However, there are thing we cannot easily do by alignment. Placing two lenses at an arbitrary distance away from one another is a difficult thing. There are a multitude of tricks we use to align components, one of them is to rely on the fact that if a beam is decently focused in the back focal plane of a lens, it comes our collimated on the other side. It is relatively easy to check that a beam is collimated by looking at how it propagates over long distances. If the beam diameter changes a lot as it propagates, then it is not collimated.

In the case of the Ti2, you will have to rely on alignment as it is unlikely that we will get the internal drawing of this microscope. Therefore, it is important to come up with a strategy for alignment. In your case, you write that you want a 70-um diameter illumination field. Assuming you use a “regular” scientific camera, this illumination field should be fairly visible in the field of view of the camera. As such, you could perhaps measure it from images acquired by the microscope.

Next is the question of how to generate such an illumination field. You write that you want critical illumination. I am not going to question this choice. In other words, I think what you want is an image of the fiber core, scaled, such that its diameter is 70um. If you were to put a 200-mm lens in the back port in a 4f configuration (what I did actually), then you would be in the same case as your detection path and for an illumination field of 70um, you would have to have an intermediate image of your core that is 100X larger, or 7000um = 7mm. Why am I suggesting this? It is simply for ease of alignment. When you send a collimated beam in a 4f system, it comes out collimated, if the 4f is aligned correctly, and as I said before, it is relatively easy to check collimation.

Now the next question is how to expand your original 400-um core diameter to 7mm. There you will need to expand by 7000/400 = 17.5X. At this point, you are getting out of the Ti2 and you have more control over the optomechanics. You could try to expand with a single lens, maybe a 20X finite conjugate objective. For example, if you take this lens (I am not affiliated with MKS Newport):

MEETOPTICS | Search & Compare Quality Optics

This lens will create a 20X image of your core, so 400x20 = 8000um = 8mm diameter. This image is then demagnified by the tube lens and objective lens (your 100X) by a factor 100, leading to a diameter of 8000/100 = 80um.

Keep in mind, these are back-of-the-envelope calculation. Best would be to perform further simulations.

Take care,

Omni

[JonD]

| Illumination Method | Critical epi illumination with relay (not Kohler). |
| Optical Setup | 12.5 mm collimator (L1), 200 mm L2, 180 mm L3. |

Using such a relay for critical illumination will form an intermediate image between L2 and L3. The spacing between L2 and L3 is important to get correct. However the spacing between L1 and L2 and between L3 and L4 is essentially collimated space and can be flexible as long as it is short enough to avoid vignetting and as long as you don’t care about the angle of the rays forming the images (only important in special cases because neither cameras nor fluorophores are very sensitive to angle of light).

The intermediate image size in the optics you propose is 0.4mm*200/12.5 = 6.4mm and the NA is modest too so I really doubt vignetting will be a problem.

So I would say in this optical configuration you don’t need to worry about the exact optical spacing between the 180mm lens and the objective.

I briefly looked at the paper you linked but it seems this paper has an additional lens “WFL” in Fig1a that you didn’t mention. My earlier description applies to what I initially understood as directly imaging the fiber tip onto the sample, not what the paper describes.

[Omnistic]

@JonD

The paper was from me I believe, it is not what @Yossi_Steinberger wants to do. They had just previously asked me to describe my system in more detail. Also as side note, after contacting the authors they explained that WFL is a 200-mm focal length lens (in case you are interested there are a few other things they didn’t disclose in the paper, just send me a message if anyone is interested, although once more, it isn’t the topic of this thread!).

Take care,

Omni