You could test it using a standard curve using some positive control. Set up a dilution series of the test DNA and measure the amplification efficiency.. You can work out the amplification efficiency by comparing the expected Ct number between standards and the actual Ct between standards  (ie. for a log2 standard curve, you would expect 1 Ct between standards. For a log10 standard curve, you would expect log_2(10)=~3.32 Ct between standards). If your actual Ct/expected Ct between standards is very close to 1, then your assay is good. 

Another way to put this is if you construct a curve from a ten-fold dilution series, the slope of the line generated from plotting the log_10(amount of standard) vs. Ct should give a slope of -1/3.32 = -0.3. The closer your line is to this, the better your assay is.

Dear Ikram

the reason for keeping your product short (i.e < 200bp) is to keep your primer efficiency high. This is an absolute pre requisite for qPCR.

In the past I have noticed that you can  - with well designed and well optimised primers - go as high as (say) 350bp and still have 'respectable' efficiencies; In my case ~ 75-85%

So my answer would be go ahead and try amplifying something of 350bp if absolutely necessary but perhaps do the following (if not already):

1. Pre screen your primers by gel based RT PCR: For products < 500bp a 2% gel is routinely used and try an annealing gradient of TM-3C to Tm+2C in 1C intervals: Pick the annealing temp which gives you the strongest (i.e. most efficient) specific band. For a product of 350bp I would veer towards Tm-2/3C to keep efficiency high

2. Given that for a product of that size you efficiency from a standard curve will probably approximate to 75% to 85% I would recommend calculating Relative quantities of mRNA using the Pfaffl method, where the actual efficiency is taken into account; rather than the standard Livak or delta delta Ct (Cq) method where the calculation presupposes a doubling of '2' i.e. perfect exponential appreciation of product

3. I will also add calculating the Tm of your primers in the context of 'real' reaction conditions i.e. with dNTP, Mg and primer concentration included in the calculation is preferable to the standard data sheet calculations (that come with primers) based purely on primer base composition. This is because binding of your primer to your target sequence is not just based on intrinsic base composition, i.e. % GC and AT and bias through out the primer of GC and AT motifs ( so called 'nearest neighbour' effects); the concentration of ions and dNTPs'  will also effect  duplex stability and hence the real time empirical melting temp. I find this algorithm particularly useful in that regard:

http://www.idtdna.com/calc/analyser

Finally, I would also add that checking your target sequence for GC content in the amplified region is useful: if your target sequence has an average of >70% GC or consecutive (> 4 runs) of G & C, this can undermine amplification efficiency. That being so add 5% DMSO to the qPCR reaction or alternatively 0.5M betaine. This will increase amplification efficiency which is paramount for products >/= 350bp

In general keep your qPCr amplicons @ 100-200bp where possible and this will increase your chances of >85-95% primer efficiency which is ideal for accurate qPCR

One more thing as I failed to answer your question completely. If you cannot alter the position of your primer except to make the product 450bp to 500bp - which will impact on primer efficiency - then with current location considering the abundance of A & T residues you should try if at all possible to end the primer in GG; GC or CC. That will increase your priming efficiency with an AT rich primer although once again priming efficiency might be an issue

Dear Laurance

Thank you so much for taking so much time out to give these suggestions. I received the primers yesterday and I will now test them on my plasmids (after diluting the plasmids) because I don't want to test them on ChIP-ed DNA right away because of its ChIP DNA is precious. I will keep the points that you mentioned in my mind.

Thank you

Ikram