We describe an improved microvolumeter (MVM) for rapidly measuring volumes of small biological samples, including live zooplankton, embryos, and small animals and organs. Portability and low cost make this instrument suitable for widespread use, including at remote field sites. Beginning with Archimedes’ principle, which states that immersing an arbitrarily shaped sample in a fluid-filled container displaces an equivalent volume, we identified procedures that maximize measurement accuracy and repeatability across a broad range of absolute volumes. Crucial steps include matching the overall configuration to the size of the sample, using reflected light to monitor fluid levels precisely, and accounting for evaporation during measurements. The resulting precision is at least 100 times higher than in previous displacement-based methods. Volumes are obtained much faster than by traditional histological or confocal methods and without shrinkage artifacts due to fixation or dehydration. Calibrations using volume standards confirmed accurate measurements of volumes as small as 0.06 µL. We validated the feasibility of evaluating soft-tissue samples by comparing volumes of freshly dissected ant brains measured with the MVM and by confocal reconstruction.
The ability to measure small volumes is important to several disciplines in biology, including physiological, evolutionary, and ecological studies of the scaling of body parts to overall body size (1-4). Comparative studies that employ volumetry often use organisms collected in remote locations, which can preclude the use of sophisticated technologies. Moreover, as further described here, current microvolumetric methods are relatively slow and expensive. Thus, we have developed a microvolumeter (MVM) and associated procedures that is portable for field use, rapid, and inexpensive.
There is a paucity of accurate volume data from small, irregularly shaped biological structures that are not easily modeled as simple geometric shapes. For example, almost all brain-body scaling studies have been limited to relatively large vertebrates (2), despite the overwhelmingly greater biodiversity of much smaller invertebrates (5). The shortage of individual small-volume data reflects both practical and cost limitations. Conventional methods of measuring small brain sizes involve measuring the mass, requiring an expensive and delicate microbalance (6). Reconstructing estimated volumes from histological sections (7) is expensive, time-consuming, and subject to shrinkage artifacts. In studies of zooplankton body volumes, due to equipment limitations, individuals are grouped prior to measurements to obtain a mean individual value (8). The techniques presented here eliminate these difficulties and provide a means of calibrating other methods that are prone to artifacts.
Our method is based on Archimedes’ principle; namely, that adding an arbitrarily shaped object to a fluid-filled container displaces an equivalent fluid volume. Previous displacement-based devices have relied upon observations of a meniscus level within a capillary, which is imprecise for small volumes. The descriptions of prior methods (9-11) lacked detailed evidence pertaining to accuracy or precision, but all fall far short of the accuracy demonstrated here. Ciborowski (10) described a simple U-shaped apparatus, consisting of an open-ended receptacle connected to a vertically oriented capillary for monitoring the meniscus level. Volumes as low as ~5 µL were measured, but because 70% ethanol was needed to reduce surface tension in the capillary, neither live specimens nor fresh tissues could be examined. Berardi (9) previously claimed accurate measurements down to 10 µL with a similar device, but provided no supporting data. Another design (11) incorporated a Gilmont micrometer syringe for adjusting the fluid level, and the device was suitable for living specimens. Although few details were provided concerning accuracy, for a 6.8-µL standard volume, a standard error of ±15% was observed. The present MVM is much more reliable, even when measuring volumes 100× smaller. We combined several strategies to minimize measurement errors, and demonstrated the high levels of accuracy and precision that can be achieved with these techniques.Materials and methods
Different versions of the same basic design were developed to encompass a large range of potential sample volumes, ranging from 20 mL (corresponding to whole bodies of some large insects) to 0.06 µL (brains of some ants and small bees). Each volumeter configuration was tested by measuring volumes (V) of spherical reference standards of known diameters and comparing measured with expected volumes according to V =(4/3) πr3, where r is a sphere's radius. The reference standards were all water-impermeable and resistant to corrosion in water or salt solutions: glass marbles (r =5 mm), stainless steel ball bearings (r =3.2, 1.5, 1.0, and 0.50 mm), and synthetic ruby balls (r =0.39, 0.30, and 0.25 mm; Small Parts, Inc., Miramar, FL). Nominal ball diameters were confirmed with a digital micrometer or from photomicrographs of the three smallest ball sizes (n ≥ 3 balls at each size). All but two deviations measured from expected diameters were ≤0.45%, and the maximum deviation was 1.4% for a 0.50-mm ball. Since such small deviations may be largely attributable to our own measurement error, we simply used nominal diameters to calculate expected ball volumes.
For larger volumes (~1–20 mL), standard disposable syringes with the plungers removed served as sample receptacles, with a smaller adjusting syringe connected via 1/4-in polypropylene tubing (Figure 1A). Typically, relatively large sample volumes are measured simply by noting a change in the initial fluid level when a sample is added to a fluid-filled receptacle. Improved precision results from attaching a syringe to the receptacle and adjusting the fluid height to a fixed criterion level (Figure 1, A and B). Thus, the sample volume is the difference between the syringe plunger's positions before and after introducing the sample. Other improvements are to produce a more consistent meniscus shape by first pushing the fluid up a short distance, then bringing it down to the criterion level, and to repeat measurements both before and after adding a sample. All of our measurements used these basic techniques.