Contact vs Noncontact Printing

Like DNA microarray fabrication, protein microarray fabrication is dependent on applying the protein onto the substrate reliably and reproducibly. To achieve this objective, two major classes of spotting, or microarray printing, have evolved. Noncontact printing is the mechanism of spotting onto the slide or plate surface without any contact. The volumes range from picoliters up to hundreds of microliters having coefficients of variation (CVs) of less than 10%, depending on the technology used. One such technology, solenoid-based dispensing systems, relies on the rapid opening and closing of valves, which permits the flow of pressurized liquid. Rapid opening and closing of a valve, or pulsing, results in a very small volume being deposited. Leaving the valve open longer results in the deposition of larger volumes. Solenoid-based systems typically spot between 10 nl and hundreds of microliters. Another noncontact technology, piezoelectric-based dispensing systems, can dispense as little as picoliter quantities of liquid. They rely on electrical pulses that create a pressure wave when a piezoelectric element is deformed and squeezes the tubing, the pressure created results in the displacement of a small volume of liquid. By opening the valve at the precise moment that the piezoelectric element is deformed, a small volume of liquid is deposited on the substrate.

The main drawback of noncontact printing is that it is not the ideal tool for arraying more viscous or concentrated protein mixtures. Cell lysates immediately come to mind when considering heterogeneous materials that may cause problems for noncontact arrayers. There is a high potential for clogging not only the dispensing solenoids, but the tubing as well. Another drawback is that the noncontact dispensing robots available are limited by the number of valves that can be used to array spots onto a slide or plate; thus, throughput becomes a problem. When printing thousands of different samples, the number of solenoids becomes the limiting step.

Contact printers address these shortcomings directly. Although there are numerous variants of contact printing technologies, the two most prevalent are the solid pin and split pin methods. Unlike the solenoid- and valve-based systems, pin-based printing systems are not limited by size. Some printers use the traditional 9- mm pin spacing used in 96- well microtiter plates, but the majority use 4.5-mm pin spacing. More recently, printers that use 2.25-mm pin spacing have become available. Depending on the size of the printhead, as many as 256 pins in an 8 x 32 configuration can be fitted when the 2.25-mm spacing is used. This results in 256 samples for every sample uptake, or over 1000 samples with only four sample uptakes. The basics are the same in solid pin and split pin printing. The robot dips the pins into the samples, the pins deposit the sample on a substrate by making contact, the pins re-dip when necessary, and the pins are washed before the next sample. Solid pins must re-dip after every deposit on a substrate; split pins act more like a quill and fill up a reservoir, of which only a fraction is deposited every time the pins make contact.

The number of spots that can be made after sample uptake by split pins varies greatly depending on a number of factors. The viscosity of the sample, the substrate type due to varying amounts of adsorption, the humidity, the pin design, and whether or not it has a reservoir are all factors and must be tested for empirically. When making DNA arrays, the samples are more homogeneous; they have similar viscosities, similar concentrations (moles of DNA), and similar molecule substrate interactions. When dealing with protein micro-arrays these characteristics are seldom the same. Proteins are usually dialyzed after purification with whichever buffer will keep the protein in solution. This can very widely from protein to protein. This will contribute to potential viscosity differences. Protein concentrations can vary greatly depending on the success of expression and subsequent purification of the protein. In case the concentrations are similar, proteins vary in size and there will be a heterogeneous distribution of the number of molecules per spot. The amino acid sequence of the protein(s) will also influence how it interacts with the substrate on which it is being spotted. In addition, the isoelectric point of the proteins will undoubtedly have an effect on how they bind to the substrate, and furthermore, hydrophilic proteins will behave differently than hydrophobic proteins. There is also the problem of protein aggregation, or too high a concentration, clogging a split pin. There are various-sized split pins with chan nels up to 400 |im across which would accommodate most substances. As attempts to make higher-density arrays to increase throughput take place, this option become less feasible.

When the substrate to be arrayed is not compatible with the split pins necessary to meet the array density required, solid pins must be employed. The need for solid pins is more of an issue when arraying cell lysates, cells of bacterial culture fractions. Solid pins are a better choice for these situations, but at the expense of throughput. Even the fastest solid pin machine, the Aushon 2470, requires twice the number of pins to outperform the fastest split pin systems, such as the Omnigrid line of microarray printers manufactured by Genomic Solutions. Throughput may or may not be an issue, but when dealing with proteins, most researchers need the process to take as little time as possible.

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