Figure 1: Binding of anti-rabbit antibodies to rabbit proteins measured in our device. A: The first picture of the measurement. B: The last picture of the measurement. C: The quotient picture of the first and last picture. D: Schematic illustration of the spot pattern on the slide
In order to test our device we immobilized rabbit-derived proteins on an iRIf slide in distinct spots (figure 1 D). These proteins were polyclonal antibodies against HCV (Hepatitis C Virus) raised in rabbit. They were detected using anti-rabbit antibodies that bind specifically to the constant regions of all rabbid derived antibodies. We used them as interaction partners in a series of experiments to reproduce a measurement previously performed in the professional device.
The binding layer on the iRIf slide consisted of an APTES/PDITC surface. The spots on the slide were produced by pipetting 3 µl [500 µg/ml] rabbit anti-HCV protein and 3 µl [1 mg/ml] BSA in an alternating pattern onto the slide. After incubation o/n the slide was blocked for 30 min with BSA [10 mg/ml].
Figure 2: Picture of our first iRIf device prototype. Note that the Nikon camera was replaced with a Canon 50D.
The camera used for the measurement was a Canon 50D. The camera was set to automatically acquire pictures at an interval of five seconds. Before the solution was flushed over the chip, the exposure time was set to obtain apprx. 80% saturation (Fig. 1 A & B).
The antibody solution was pipetted into the flow chamber without the use of a microfluidic pump. Instead, a syringe was loaded with 500 µl [5 µg/ml] anti-rabbit antibody solution (diluted in PBS) and slowly released into the binding chamber of the device by gently dispensing it from the syringe.
As can be seen in figure 1 C, the quotient picture clearly shows binding of anti-rabbit antibodies to the rabbit protein spots. The BSA control spots show none or negligible unspecific binding.
Spot analysis by hand: A step by step guidance
This step by step guidance shows you how to dertermine endpoint intensities of the binding experiment as shown above. Have a look at our iRIF page to getall the background informations about the detection method.
- During your iRIF experiment you take two pictures. One before the binding event and one after the binding event. Open both pictures in ImageJ.
- Go to "Process" and click on "Image Calculator..."
- Set "Image1" to the image after the binding event and "Image2" to the image before the binding event. Choose "divide" as operation. Activate the "32-bit (float) result" and click "ok". A quotient picture will be calculated and opened.
- Go to "Image" then "Adjust" and klick on "Brightness/Contrast..." and play around with "brightness" and "contrast" till the spots get visible.
- Mark the spots using the "oval" selection tool and press "T" on your keybord. Do this for every spot. Do not foret to make a selection of the background.
- Go to the ROI-manager that poped up when you first pushed T and click on "Deselect" and then on "Measure". The results of your Measurment will pop up. You can save them in Excel.
Remember that signals will only differ by about 0.1 - 0.8 percent depending on how strong your binding is.
How To Build Your Own Device
In this section we demonstrate how to build your own iRIf device from scratch, using affordable, low-tech material. The design is focused on creating a device that is both low-priced and portable, demonstrating the potential for the DiaCHIP to be used even in areas where high-tech laboratories are inaccessible.
General Principle
Figure 3: Our device from top.
As seen in figure 3, the basic setup is fairly simple. Light from an LED enters a lens to obtain a parallel light beam. To achieve this, the distance from the LED to the lens should be close to one focal length.
Next, the light hits the iRIf slide where it is reflected (in the same angle as it hits the slide) and enters a second lens that projects an image of the slide on the CCD chip of the camera.
Construction Guidance
To rebuild our iRIf device, we used the following parts:
| Part | Description | Manufacturer / Part name | Price[USD] |
| A fairly strong LED | In our case a 3W green 520 nm LED | Cree XP-E on star circuit board | 10 |
| A cooling element | Used to prevent the LED from overheating | Fischer Elektronik ICK S | 9 |
| A constant-current-source | To ensure static LED light (no flickering) | Roschwege GmbH KSQ-3W | 15 |
| An AC/DC rectifier | Used to grant direct current to the LED | Goobay 67951 | 16 |
| Two optical lenses | Here we used two identical 60 mm lenses | Thorlabs AC254-060-A | 160 |
| A camera | We used a SLR camera | Canon 550D | 200-400 |
| A microfluidics chamber | Our chamber consists of an iRIf slide attached to a PDMS flow cell | Made ourselves | |
| Magnets | Used to attach the flow cell to the device | 5 mm Neodymium magnets | 3 |
| A small syringe | We used a regular 10 ml syringe | BRAUN INJEKT 10 ml | < 1 |
| A microfluidic tube | Used to connect the flow chamber with the syringe | - | < 1 |
| A case for the device | 5 mm thick acrylic glass cutouts to hold the parts in place | Ordered at http://formulor.com | 100 |
Figure 3: (A) The LED glued to the cooling element with thermal adhesive; (B) The 10 magnets, each 5x5x5 mm as well as a PDMS flow chamber (depth of the flow chamber: 30 µm); (C) One of the lenses used in the setup
A major problem that we encountered when building the device from scratch was to ensure that all components are at the proper distance and angle to each other. Correct alignment is crucial as a slight misplacement of a component may lead to lower signal strength, blurred images or, in the worst case, no signal at all. This can be difficult since our device does not rely on straight angles. We overcame this challenge by designing a case for the device that ensures the right placement of the components inside the device. We calculated all distances between the LED, lenses, camera and slide using the laws of geometrical optics and drew an vector graphic blueprint for our device. Next, we constructed a digital 3D model of the casing based on the vector blueprint to avoid an expensive and time-consuming trial and error process (Figure 4).
Figure 4: 3D model of our device, built from the vector files used to order the parts.
Figure 5: The 3D model of our device without walls and top part. Parts have been colorized for clarification - Green: The wall where light exits the device and enters the camera; Pink: a platform holding the cooling element and LED in place; Blue: platforms for holding the lenses in place; Gray: rear end wall where the flow chamber is attached to. The slits in the top and bottom part where the magnets need to be attached.
The casing is designed to keep all the necessary parts (lenses, LED) in place safely during a measurement, but allows them to be removed to facilitate transportation of the device (i.e. to the Giant Jamboree). Figure 5 illustrates the parts that hold the lenses and LED in place.
After setup of the 3D model of our device, we created a vector graphic of all required parts in a 2D plane. Using this vector graphic, we ordered the parts at Formulor, a service which uses laser cutting to cut out parts from acrylic glass and other materials. The cutting laser is guided by the paths defined in the vector graphic template. The template is shown in figure 6, and may be downloaded and used by everyone to build their own device. To allow easy mounting of the casing, we also provide an easy to understand manual. One advantage of acrylic glass is that the parts can be glued together easily using a few drops of acetone. The vector graphic also contains a template for the parts which are necessary to attach the glass and PDMS slide to the device. They should be cut from 1 mm thick acrylic glass. The microfluidic tubes can be glued to these parts (figure 7) and attached to the device with the magnets (figure 8)
Figure 6: An image of the vector file used to order the parts. The vector file is used to excise parts of a 5 mm acrylic glass board with a laser. Black lines are cut out by the laser, gray lines/areas are engravings. Refer to the
download section to downloaded the vector graphic files.
The most difficult part to build is the PDMS flow chamber since it requires a microstructured silicon wafer, which can only be produced in a cleanroom environment. The template used for our PDMS flow cell is shown in figure 9. If your facility has an engineering department, there is a high chance that a cleanroom is present. Ask the personnel in charge if they can help you out with producing your flow cell. If you already use another type of microfluidic flow chamber, our device should still be compatible with it, though some adaptions might be necessary. If you have no possibility of creating a flow chamber, you might want to consider building a very simplified DIY flow chambers from two glass slides, separated by thin tape. Note that our device was not primarily build to support such a chamber and would need appropriate modifications.
Figure 7 - left: a flow chamber consisting of an iRIf slide attached to a PDMS flow cell - center: a syringe attached to a pipette tip, used to inject antibody solutions into the flow chamber - right: the part of our device which connects the flow chamber, the syringe and the iRIf device.
Figure 8: The device with and without the mounted flow chamber.
Figure 9: Layout used to produce the wafer of our PDMS flow cell.
Download section
Click on the image to download a manual which will show you how to mount the casing of the device.
Legal notice
The iRIf detection method is patented.
Biametrics
and associated persons own patents concerning the detection principle.
These patents are:
