Faq's

When starting AutoResp™ and this error message appear, AutoResp™ was not installed with administrator rights. During the AutoResp™ installation process, a file must be copied into a Windows system folder. This requires an administrator login. Here is the instruction how to install the necessary file to get rid of the error message:

1) Login as an administrator

2) Download the following configuration file, LINK, and save the file to your desktop.

3) Locate the folder folder: C:\ProgramData\Measurement Computing\DAQ\ . The folder Program Data is a system hidden folder. So you might have to change settings for the folder in order see hidden folders and system files. Hidden folders are shown as “transparent” in Windows’ file explorer.

4) If the folder doesnt exist, please create it.

5) Now move the file into this folder.

6) After you moved the file, change folder settings again to hide hidden and system files.

7) Start AutoResp™. The error message should be gone by now.

If the display on your OXY-REG is giving the error message "SE.BR" (sensor breakage), it means that the signal input type for the instrument has been changed from the default setting (potmeter).

To change the input type back to the default setting, press the OK button once. Then press the arrow buttons until the input type POTM (potmeter) is shown in the display and then press OK to choose this. Finally exit the set up menu by pressing the OK button several times until the display shows - - - -.

Tracking requires two synchronized (in time) cameras to track animals in three dimensions.

Mount the two cameras with a 90° angle in between, e.g. one camera filming from above and one filming from the side.

Simultaneous video recordings from both cameras can then be analyzed in LoliTrack to get two pairs of X-Y coordinates.

In this way, one pair of X-Y coordinates translates into the third (Z) coordinate. See the figure below.

This guide shows you how to draw zones and mask in LoliTrack and ShuttleSoft.

Use the Circle button, then draw a circle in one of the compartments. Afterwards, click on the Mask Inside Area button. 

Use the Circle button again to draw a circle in the other compartment. Afterwards, click on the Mask Inside Area button.

Use the Rectangular button, then draw a rectangle between the two compartments. Afterwards, click on the Mask Inside Area button.  

Now invert the mask by clicking on the Invert Mask button. 

Draw a rectangular area in one compartment, and click on a Zone button (A, B, C...). This area is now defined as a zone of interest. 

Draw a rectangular area in the other compartment, and click on a different Zone button. This area is now also defined as a zone of interest. 

This is an example of how a typical mask and zones could look like running a ShuttleSoft experiment.

To change the language in the PIT tag reader software (APR-PC-DEMO) from German to English, please go to the following program files folder on your PC:

1. Locate C:\Program Files (x86)\AgriDent
2. Delete the file(s) that has the *.lng file extension (e.g. Agrident.lng).
3. Without this language file, the software will now use the default language (English).

Loligo^® Systems offers the three main types of oxygen sensors differing in measuring principle, response time, sensor size, maintenance requirement and pricing.

1. Optical oxygen sensors
For many invasive techniques, measurements in tiny volumes (e.g. micro respirometry) or applications which require high temporal and spatial resolution, fibre-optic oxygen sensors are the only solution.

Even for general purposes we recommend optical oxygen sensors featuring low maintenance requirements, high stability and accuracy, no electrical interference, no ground loop problems, and zero O₂ consumption by the sensor.

The main disadvantages is price, single-channel meters start at about €3750. A high temperature sensitivity of this technology and a fragile tip on the <50-140 nm micro sensors might also be a problem in some applications. The macro sensors, on the other hand, are very tough.

2. Galvanic cell oxygen electrodes
Galvanic type oxygen probes are inexpensive and rugged sensors producing a milliVolt signal for easy instrumentation without supplying power. 

We recommend galvanic probes for applications like respirometry and field measurements, and as a low-budget alternative to optical oxygen equipment, e.g. for multi-channel systems. They can be used with relatively long cables (+25 meters). We recommend using a galvanic isolation preamplifier between the probe and any data acquisition system to minimize possible ground loop problems.

The main disadvantage of galvanic probes is a relative long response time and oxygen self-consumption making them unsuitable for measurements in small volumes and in un-mixed samples.

3. Polarographic oxygen electrodes (or Clark type electrodes)

Low oxygen self-consumption and a small size make these sensors suitable for physiological setups like blood gas analysis and low volume respirometry. 

However, Clark type electrodes require much maintenance, often membranes and electrolyte fluid should be changed on a daily basis, and polarographic amplifiers for these sensors are quite costly. 

Thus, we recommend the E101 as a replacement oxygen electrode for customers with their own polarograhic oxygen meter, e.g. PHM meter from Radiometer Copenhagen, OM200 from Cameron Instr. Co. etc.

CALIBRATION PRINCIPLE
Instrument calibration is an essential first step in analytical and measurement procedures.  The rationale for a two point calibration is to provide two points of reference with which to calibrate the instrument and, therefore, ensuring accurate measurements with your sensor of the unknown concentrations of your samples.  A two point calibration between two extremes are advocated, e.g. 0 - 100 % air saturation.  For the low calibration point, we recommend using either nitrogen gas or sodium sulphite solution, for the high calibration point, bubble atmospheric air using an aerator.  Please see below for general step-wise instructions on how to carry out a two point calibration.  For specific instructions for a particular oxygen instrument, please click on the specific links.

GENERAL

1. Connect the probe to the oxygen instrument and turn on the instrument.
2. Place the tip of the probe in a mixed air-equilibrated water sample. This can be achived by purging atmospheric air into sample water, e.g. with an air pump and air stone.
3. Wait for the reading to stabilize. Then save this point as the HIGH calibration value (100% air saturation).
4. Place the tip of the probe in an oxygen free water sample (e.g. by purging nitrogen gas into sample water or by dissolving ~10 grams of Na₂SO₃ in 500 mL of distilled water).
5. Wait for the reading to stabilize. Then save this point as the LOW calibration value (0% air saturation).
6. Rinse the probe with distilled water.
7. Place the probe into the sample, and record the sample oxygen saturation when the reading is stable.

See below for a diagrammatic representation of sensor readings at:
a)  100 %  to 0 %  air saturation
b)  0% to 100 air saturation
c)  100 % to 0 % and back up to 100 % air saturation.

100% to 0% air saturation	0% to 100% air saturation
100% to 0% to 100 % air saturation

The oxygen sensor spot should be integrated into a dry, clean vessel (humidity or oily residues affect the adhesion and may lead to a detaching of the sensor spot from the vessel wall). The adhesion surface area of the vessel wall should be plane or slightly arched but not strongly curved or corrugated. Otherwise, the spot may detach partly from the vessel wall during the curing process of the silicon glue due to the tension of the spot.

The silicon glue should be cured for at least 60 min. before the test vessel is fill.

Apply a small amount of silicon glue (D5-Spot: ca. 4 µL) onto the red side of the spot.


Position the spot on the plane bent end of a spatula.


Attach the spot to the inside wall of the test vessel.


Shift the sensor spot slightly with little pressure onto the surface so that a small ring of silicon glue emerges around its edge.


No air should be enclosed in the silicon layer between spot and vessel wall.


Silicon stains on the black sensor surface should be avoided since they increase the response time of the sensor.

The light from the blue-green LED excites the oxygen sensor to emit fluorescence. If the sensor spot encounters an oxygen molecule, the excess energy is transferred to the oxygen molecule in a non-radiative transfer, decreasing or quenching the fluorescence signal. The degree of quenching correlates to the partial pressure of oxygen in the matrix, which is in dynamic equilibrium with oxygen in the sample. The decay time measurement is internally referenced.

The Loligo^® shuttle tanks are modified versions of the classic operant conditioning chambers (also known as the Skinner box) used for experimental analysis of behavior, e.g. to study operant conditioning and classical conditioning in animals.

An operant conditioning chamber permits experimenters to study behavior conditioning (training) by teaching a subject animal to perform certain actions (like pressing a lever) in response to specific stimuli like a light or sound signal. When the subject correctly performs the behavior, a mechanism delivers food or another reward. In some cases, the mechanism delivers a punishment for incorrect or missing responses.

With this apparatus, experimenters perform studies in conditioning and training through reward/punishment mechanisms. Operant chambers have at least one operandum, that can automatically detect the occurrence of a behavioral response or action.

Typical operanda for primates and rats are response levers. Despite such a simple configuration (e.g. one operandum and one feeder), it is possible to investigate many psychological phenomena in this way.

For this reason, operant conditioning chambers have become common in a variety of research disciplines including behavioral pharmacology, and Skinner's Box have been used extensively for behavioral research in primates and rats.

Loligo^® shuttle tanks have been developed for aquatic animals, like zebrafish or crustaceans, and the tank design allows for independent control of water quality in two sub compartments. Tank dimensions are made special to accomodate a wide variaty of animal species and sizes.

Inside the Shuttle tank, the animal can freely "shuttle" between two sub compartments with opposite acting controls.

The computerized Loligo^® shuttle systems are equiped with a video camera and lighting conditions enabling real-time pc vision software to detect animal locomotion.

If the animal changes its position from one user-defined zone to the next through locomotion, the computer software (ShuttleSoft) activates/deactivates programmed devices to change environmental conditions inside the tank, e.g. to regulate water temperature to preferred values through behavior. Or you can set up two different (constant) temperature levels in the two tank compartments independent of fish behavior for exposure/avoidance/choice tests.

Today, a main application of Loligo^® shuttle tanks measurements of temperature preference in aquatic ectotherms (as well as avoidance behavior), and automated computerized systems, have been made for a range of other environmental factors like water turbidity, salinity, oxygen saturation, pH and pCO2.

The turnkey systems offered include everything needed for video behavior analysis as well as monitoring and regulating water quality.

Intermittent flow (or stop-flow) respirometry - Principles and Background

Measurements of oxygen consumption rates on fish and other water breathers commonly involves using one of three different methods:
 
1. Closed respirometry (or constant volume respirometry) 
2. Flow-through respirometry (or open respirometry) 
3. Intermittent flow respirometry (or stop-flow)

1. Closed respirometry (or constant volume respirometry)

Measurements in a sealed chamber of known volume (a closed respirometer). The oxygen content of the water is measured initially (t0), then the respirometer is closed and at the end of the experiment (t1) the oxygen content is measured again. 
Knowing the body weight of the animal, the respirometer volume and the oxygen content of the water at time t0 and t1, the mass specific oxygen consumption rate (mg O₂/kg/hour) can be calculated as follows:

VO₂ = ([O₂]t0 – [O₂]t1) • V/t/BW

• [O₂]t0 = oxygen concentration at time t0 (mg O₂/liter)
• [O₂]t1 = oxygen concentration at time t1 (mg O₂/liter)
• V = respirometer volume minus volume of experimental animal (liter)
• t = t1 – t0 (hour)
• BW = body weight of experimental animal (kg)

An advantage of this method is its simplicity. A disadvantage is that the measurements are never made at a constant oxygen level, due to the continuous use of oxygen by the animal inside the respirometer. This might cause problems when interpreting data, since animal respiration often changes with ambient oxygen partial pressure.
Furthermore, metabolites from the experimental animal, i.e. CO₂, accumulate in the water, thus limiting the duration of measurements. This limited time for measurements prevents the experimental animal to recover from initial handling stress that often increase fish respiration significantly and for several hours, thus overestimating oxygen consumption rates.

2. Flow-through respirometry (or open respirometry)
This is a more sophisticated method for oxygen consumption measurements. Experimental animals are placed in a flow-through chamber with known flow rate. Oxygen is measured in the inflow and outflow and oxygen consumption rate (mg O₂/kg/hour) can be calculated as:

VO₂ = F • ([O₂]_in – [O₂]_out)/BW

• F = water flow rate (l/hour)
• [O₂]_in = oxygen content in water inflow (mg O₂/liter)
• [O₂]_out = oxygen content in water outflow (mg O₂/liter)
• BW = body weight of experimental animal (kg)

The advantages of this method are several:

1. The duration of the experiment is in principle unlimited
2. No accumulation of CO₂ and other metabolites
3. Its possible to measure at a constant oxygen level
4. By controlling the quality of the inflowing water its possible to measure metabolism at different desired levels of oxygen, salinity etc.

However, this method bring along one significant disadvantage: In order to determine oxygen consumption by open respirometry it is crucial that the system is in steady state. This means that the oxygen content of the in-flowing and out-flowing water, AND the oxygen consumption of the animal, have to be constant. 
If the oxygen consumption of the animal for some reason changes during the experiment, steady state will not exist for a while. The above formula will not give the correct oxygen consumption rate until the system is in steady state again. The duration of the time lag depends on the relationship between chamber volume and flow rate. Thus, open respirometry measurements have poor time resolution and are not suitable for determination of oxygen consumption on organisms with a highly variable respiration like fish.

3. Intermittent flow respirometry
Our system for automatic respirometry works by intermittent flow respirometry aiming at combining the best of both 1) closed and 2) flow-through respirometry.

• The experimental animal is placed in a respirometer immersed in an ambient tank (temperature bath).
• A recirculating pump ensures proper mixing of the water inside the respirometer and adequate flow past the oxygen probe. 
• A second pump can swap the water inside the respirometer with water from the ambient tank (temperature bath). During measurements of oxygen consumption, this flush pump is turned off and the systems operates like a closed respirometry setup. 
• Then the pc controlled flush pump turns on pumping ambient water into the respirometer and bringing the oxygen content back to pre measurement levels.
  In this way, problems with accumulating metabolites and severe changes in oxygen level due to animal respiration are avoided.
• As with open respirometry, the duration of the experiment is in principle unlimited.

However, the most important advantage is the great time resolution of this method. Oxygen consumption rates of animals can be determined for every 10th minutes over periods of hours or days, making our systems for automatic respirometry extremely suited for uncovering short term variations (minutes) in respiration.

In summary, our system for automatic respirometry is developed for prolonged and automatic measurements of oxygen consumption rate in a controlled laboratory environment.

The automatic measuring proceedure runs in 3 phases:

1. Measuring period
2. Flush period
3. Wait period

In the Measuring period (1) the flush pump is off, and the chamber is closed. Fish respiration rate is calculated from the decline in oxygen. During this time the recirculation pump is active to mix the water inside the respirometer and to ensure proper flow past the oxygen sensor.
The measuring period is followed by a Flush period (2) where the flush pump is active pumping water from the ambient temperature bath and into the respirometer. During this period the recirculation pump is inactive and the oxygen curve will raise to approach the level of the amient water.
Finally, the flush pump stops and the loop ends with a short Wait period (3) before starting a new measuring period. This waiting period is necessary to account for a lag in the system response resulting in a non-linear oxygen curve. During the Wait period the recirculation pump is active.

Examples of raw MO₂ data

Standard metabolic rate of juvenile Rainbow Trout was determined in static respirometer chambers and with an automated respirometry system from Loligo^® Systems. Initial high oxygen consumption rates due to handling stress, were followed by a gradual decline to lower and more stable values indicating standard metabolic rate for the specimen. Notice the high temporal resolution (10 min) of the system revealing sudden changes in MO₂.

Courtesy by J.Svendsen/DIFRES & J.Lund/Univ. Aarhus

It is important that the volume and dimensions of a respirometer chamber fits the body dimensions and respiration rate of the experimental animal.

Temperature affects metabolic rate in ectotherms (like fish). High experimental temperatures thus favor the use of larger chambers and visa versa.

Metabolic rate (or respiration) also scales with the body size of the animals, e.g. large animals have lower mass specific O₂ cosumption rates than smaller animals (e.g. mouse-and-elephant example).

If the chamber volume is too large, the resulting oxygen depletion curve will be too flat for reliable estimates of the slope that is used in the calculation of the oxygen consumption rate (MO2). A chamber much larger than the animal will allow space for unwanted activity if trying to measure standard metabolic rate of an in-active (static or resting) animal.

On the other hand, a large fish in a small chamber means that oxygen will decline rapidly to hypoxic levels which might affect the metabolism of oxyregulatory species.

As a rule of thumb, we recommend a chamber volume some 10-50 times the volume (≈ wet weight) of the animal depending on the temperature, the species and the size of the specimen, e.g. a 5 L chamber for a large adult 500 g fish at low temperatures (ratio 1:10) or a 500 ml chamber for a small 10 g juvenile fish at higher temperatures (ratio 1:50).

Active fish swimming in a tunnel respirometer have high mass-specific oxygen consumption rates (MO2), allowing reliable oxygen consumption estimates in relative higher respirometer volume than for static respirometry. Tunnel respirometers are difficult to build with a fish volume to respirometer volume of less than 1:100. We recommend a maximum ratio between the wet weigth (or volume) of the fish and the volume of the swim respirometer of 1:200, e.g. a 90 liter swim respirometer fits fish down to app. ½ kg. If the volume is too large, the resulting oxygen curve will be too flat for reliable estimates of the slope that is used in the calculation of oxygen consumption rate (MO2).

Another restraint is the dimensions of the test section, which should allow the experimental fish to perform unrestricted swimming. This will largely depend on the species and mode of swimming.

Please contact us to get free advice on which swim tunnel that will be best for your project.

To wash out a tube-shaped respirometer chamber requires a volume ~5 times that of the chamber.

Thus, if you use a pump delivering a flow per minute equal to the chamber volume, then a 5 minutes flush period will ensure that 99% of the water is replaced during each flushing cycle.

Stronger pumps might create too strong a flow forcing fish to struggle or unwanted activity. Lower flows require longer time for flushing, and this will decrease time resolution of MO2 data.

Some materials, like Plexiglass/Perspex can act as oxygen stores and sinks, forming pools of oxygen which can be released or stored in a reversible way - see Stevens, J. (1992) J. Appl. Physiol. 72, 801-804. 

This can have an important effect when chambers are down-scaled for micro respirometric measurements of very low oxygen fluxes. 

Thus, for oxygen measurements in small volumes less than a few millilitres (mL), we recommend chambers of glas and inert components with non or low oxygen storage capacity! 

For respirometers with larger volumes (>½ litre), the use of acrylic materials have no significant effects on measurements.

If you purchased a software upgrade, please follow the steps below to add new license codes to your WIBU dongle:

1) Download the following info file, LINK, and save the file to your desktop.

2) Double-click it to transfer licens codes from your dongle (connected to your PC) and save it to a .wbc file.

3) Email the .wbc file to this address: mail@loligosystems.com.

4) We will then quickly return a .rtu file as an email attachment. This .rtu file carries the new license code(s).

5) Save the .rtu file on your desktop.

6) Double-click the .rtu file to transfer new license code(s) to the dongle connected to your PC.

7) Finally, download the upgraded version of the software from our website and install it.